What is a primer in ecology statistics?

A Primer of Ecological Statistics emphasizes a general introduction to probability theory and provides a detailed discussion of specific designs and analyses that are typically encountered in ecology and environmental science.

What is primer software used for?

It is primarily used in the scientific community for ecological and environmental studies . Multivariate routines include: grouping (CLUSTER) sorting (MDS)

How do you choose a primer for DNA replication?

Taking into consideration the information above, primers should generally have the following properties: Length of 18-24 bases. 40-60% G/C content. Start and end with 1-2 G/C pairs. Melting temperature (Tm) of 50-60°C. Primer pairs should have a Tm within 5°C of each other. Primer pairs should not have complementary regions.

What is ecological primer?

The Primer of Ecological Restoration is a succinct introduction to the theory and practice of ecological restoration as a strategy to conserve biodiversity and ecosystems .

What is a statistics primer?

The primer document covers the fundamental principles of statistics and probability like variables, populations and samples, central tendency, variability, hypothesis testing, measures of association, comparative measures, probability basics, and more.

What is a primer in gene expression?

A primer is a short nucleic acid sequence that provides a starting point for DNA synthesis . In living organisms, primers are short strands of RNA. A primer must be synthesized by an enzyme called primase, which is a type of RNA polymerase, before DNA replication can occur.

What are the primers in genomics?

Primers are short stretches of DNA that target unique sequences and help identify a unique part of genome — let's say, a gene. Primers are usually 18 to 25 nucleotides long. They can be synthesized in a special lab, and are used in many different ways.

A Primer of Ecological Statistics - Nicholas J. Gotelli, Aaron M. Ellison 2th Edition [PDF]

A Primer of Ecological Statistics SECOND EDITION

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Estadísticos e-Books & Papers

A Primer of

Ecological Statistics Second Edition

Nicholas J. Gotelli Aaron M. Ellison

University of Vermont

Harvard Forest

Sinauer Associates, Inc. Publishers Sunderland, Massachusetts U.S.A.

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A PRIMER OF ECOLOGICAL STATISTICS, SECOND EDITION

Copyright © 2013 by Sinauer Associates, Inc. All rights reserved. This book may not be reproduced without permission of the publisher. For information address Sinauer Associates Inc., 23 Plumtree Road, Sunderland, MA 01375 U.S.A. FAX 413-549-1118 EMAIL [emailprotected], [emailprotected] WEBSITE www.sinauer.com

Library of Congress Cataloging-in-Publication Data Gotelli, Nicholas J., 1959A primer of ecological statistics / Nicholas J. Gotelli, University of Vermont, Aaron M. Ellison, Harvard University. -- Second edition. pages ; cm Includes bibliographical references and index. ISBN 978-1-60535-064-6 1. Ecology--Statistical methods. I. Ellison, Aaron M., 1960- II. Title. QH541.15.S72G68 2013 577.072--dc23 2012037594 Printed in U.S.A. 5 4 3 2 1

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For Maryanne & Elizabeth

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Who measures heaven, earth, sea, and sky Thus seeking lore or gaiety Let him beware a fool to be. SEBASTIAN BRANT, Ship of Fools, 1494. Basel, Switzerland.

[N]umbers are the words without which exact description of any natural phenomenon is impossible…. Assuredly, every objective phenomenon, of whatever kind, is quantitative as well as qualitative; and to ignore the former, or to brush it aside as inconsequential, virtually replaces objective nature by abstract toys wholly devoid of dimensions — toys that neither exist nor can be conceived to exist. E. L. MICHAEL, “Marine ecology and the coefficient of association: A plea in behalf of quantitative biology,” 1920. Journal of Ecology 8: 54–59.

[W]e now know that what we term natural laws are merely statistical truths and thus must necessarily allow for exceptions. …[W]e need the laboratory with its incisive restrictions in order to demonstrate the invariable validity of natural law. If we leave things to nature, we see a very different picture: every process is partially or totally interfered with by chance, so much so that under natural circ*mstances a course of events absolutely conforming to specific laws is almost an exception. CARL JUNG, Foreword to The I Ching or Book of Changes.Third Edition, 1950, translated by R. Wilhelm and C. F. Baynes. Bollingen Series XIX, Princeton University Press.

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Brief Contents PART I FUNDAMENTALS OF PROBABILITY AND STATISTICAL THINKING 1 2 3 4 5

An Introduction to Probability 3 Random Variables and Probability Distributions 25 Summary Statistics: Measures of Location and Spread 57 Framing and Testing Hypotheses 79 Three Frameworks for Statistical Analysis 107

PART II DESIGNING EXPERIMENTS 6 Designing Successful Field Studies 137 7 A Bestiary of Experimental and Sampling Designs 163 8 Managing and Curating Data 207

PART III DATA ANALYSIS 9 10 11 12

Regression 239 The Analysis of Variance 289 The Analysis of Categorical Data 349 The Analysis of Multivariate Data 383

PART IV ESTIMATION 13 The Measurement of Biodiversity 449 14 Detecting Populations and Estimating their Size 483 Appendix Matrix Algebra for Ecologists 523

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Contents PART I

Fundamentals of Probability and Statistical Thinking CHAPTER 1

An Introduction to Probability 3

Bayes’ Theorem 22 Summary 24

What Is Probability? 4 Measuring Probability 4 The Probability of a Single Event: Prey Capture by Carnivorous Plants 4 Estimating Probabilities by Sampling 7

Problems in the Definition of Probability 9 The Mathematics of Probability 11 Defining the Sample Space 11 Complex and Shared Events: Combining Simple Probabilities 13 Probability Calculations: Milkweeds and Caterpillars 15 Complex and Shared Events: Rules for Combining Sets 18 Conditional Probabilities 21

CHAPTER 2

Random Variables and Probability Distributions 25 Discrete Random Variables 26 Bernoulli Random Variables 26 An Example of a Bernoulli Trial 27 Many Bernoulli Trials = A Binomial Random Variable 28 The Binomial Distribution 31 Poisson Random Variables 34 An Example of a Poisson Random Variable: Distribution of a Rare Plant 36

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Contents

The Expected Value of a Discrete Random Variable 39 The Variance of a Discrete Random Variable 39

Continuous Random Variables 41 Uniform Random Variables 42 The Expected Value of a Continuous Random Variable 45 Normal Random Variables 46 Useful Properties of the Normal Distribution 48 Other Continuous Random Variables 50

The Central Limit Theorem 53 Summary 54

CHAPTER 4

Framing and Testing Hypotheses 79 Scientific Methods 80 Deduction and Induction 81 Modern-Day Induction: Bayesian Inference 84 The Hypothetico-Deductive Method 87

Testing Statistical Hypotheses 90 Statistical Hypotheses versus Scientific Hypotheses 90 Statistical Significance and P-Values 91 Errors in Hypothesis Testing 100

CHAPTER 3

Parameter Estimation and Prediction 104 Summary 105

Summary Statistics: Measures of Location and Spread 57

CHAPTER 5

Measures of Location 58 The Arithmetic Mean 58 Other Means 60 Other Measures of Location: The Median and the Mode 64 When to Use Each Measure of Location 65

Measures of Spread 66 The Variance and the Standard Deviation 66 The Standard Error of the Mean 67 Skewness, Kurtosis, and Central Moments 69 Quantiles 71 Using Measures of Spread 72

Some Philosophical Issues Surrounding Summary Statistics 73 Confidence Intervals 74 Generalized Confidence Intervals 76

Summary 78

Three Frameworks for Statistical Analysis 107 Sample Problem 107 Monte Carlo Analysis 109 Step 1: Specifying the Test Statistic 111 Step 2: Creating the Null Distribution 111 Step 3: Deciding on a One- or Two-Tailed Test 112 Step 4: Calculating the Tail Probability 114 Assumptions of the Monte Carlo Method 115 Advantages and Disadvantages of the Monte Carlo Method 115

Parametric Analysis 117 Step 1: Specifying the Test Statistic 117 Step 2: Specifying the Null Distribution 119 Step 3: Calculating the Tail Probability 119 Assumptions of the Parametric Method 120 Advantages and Disadvantages of the Parametric Method 121

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Contents

Non-Parametric Analysis: A Special Case of Monte Carlo Analysis 121

Bayesian Analysis 122 Step 1: Specifying the Hypothesis 122 Step 2: Specifying Parameters as Random Variables 125 Step 3: Specifying the Prior Probability Distribution 125

Step 4: Calculating the Likelihood 129 Step 5: Calculating the Posterior Probability Distribution 129 Step 6: Interpreting the Results 130 Assumptions of Bayesian Analysis 132 Advantages and Disadvantages of Bayesian Analysis 133

Summary 133

PART II

Designing Experiments CHAPTER 6

Designing Successful Field Studies 137 What Is the Point of the Study? 137 Are There Spatial or Temporal Differences in Variable Y? 137 What Is the Effect of Factor X on Variable Y? 138 Are the Measurements of Variable Y Consistent with the Predictions of Hypothesis H? 138 Using the Measurements of Variable Y, What Is the Best Estimate of Parameter θ in Model Z? 139

Manipulative Experiments 139 Natural Experiments 141 Snapshot versus Trajectory Experiments 143 The Problem of Temporal Dependence 144

Press versus Pulse Experiments 146 Replication 148 How Much Replication? 148 How Many Total Replicates Are Affordable? 149 The Rule of 10 150

Large-Scale Studies and Environmental Impacts 150

Ensuring Independence 151 Avoiding Confounding Factors 153 Replication and Randomization 154 Designing Effective Field Experiments and Sampling Studies 158 Are the Plots or Enclosures Large Enough to Ensure Realistic Results? 158 What Is the Grain and Extent of the Study? 158 Does the Range of Treatments or Census Categories Bracket or Span the Range of Possible Environmental Conditions? 159 Have Appropriate Controls Been Established to Ensure that Results Reflect Variation Only in the Factor of Interest? 160

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Contents

Have All Replicates Been Manipulated in the Same Way Except for the Intended Treatment Application? 160 Have Appropriate Covariates Been Measured in Each Replicate? 161

CHAPTER 8

Managing and Curating Data 207 The First Step: Managing Raw Data 208

Summary 161

Spreadsheets 208 Metadata 209

CHAPTER 7

A Bestiary of Experimental and Sampling Designs 163 Categorical versus Continuous Variables 164 Dependent and Independent Variables 165 Four Classes of Experimental Design 165 Regression Designs 166 ANOVA Designs 171 Alternatives to ANOVA: Experimental Regression 197 Tabular Designs 200 Alternatives to Tabular Designs: Proportional Designs 203

Summary 204

The Second Step: Storing and Curating the Data 210 Storage: Temporary and Archival 210 Curating the Data 211

The Third Step: Checking the Data 212 The Importance of Outliers 212 Errors 214 Missing Data 215 Detecting Outliers and Errors 215 Creating an Audit Trail 223

The Final Step: Transforming the Data 223 Data Transformations as a Cognitive Tool 224 Data Transformations because the Statistics Demand It 229 Reporting Results: Transformed or Not? 233 The Audit Trail Redux 233

Summary: The Data Management Flow Chart 235

PART III

Data Analysis CHAPTER 9

Regression 239 Defining the Straight Line and Its Two Parameters 239 Fitting Data to a Linear Model 241 Variances and Covariances 244

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Least-Squares Parameter Estimates 246 Variance Components and the Coefficient of Determination 248 Hypothesis Tests with Regression 250 The Anatomy of an ANOVA Table 251 Other Tests and Confidence Intervals 253

Assumptions of Regression 257 Diagnostic Tests For Regression 259 Plotting Residuals 259 Other Diagnostic Plots 262 The Influence Function 262

Monte Carlo and Bayesian Analyses 264 Linear Regression Using Monte Carlo Methods 264 Linear Regression Using Bayesian Methods 266

Other Kinds of Regression Analyses 268 Robust Regression 268 Quantile Regression 271 Logistic Regression 273 Non-Linear Regression 275 Multiple Regression 275 Path Analysis 279

Hypothesis Tests with ANOVA 296 Constructing F-Ratios 298 A Bestiary of ANOVA Tables 300 Randomized Block 300 Nested ANOVA 302 Two-Way ANOVA 304 ANOVA for Three-Way and n-Way Designs 308 Split-Plot ANOVA 308 Repeated Measures ANOVA 309 ANCOVA 314

Random versus Fixed Factors in ANOVA 317 Partitioning the Variance in ANOVA 322 After ANOVA: Plotting and Understanding Interaction Terms 325 Plotting Results from One-Way ANOVAs 325 Plotting Results from Two-Way ANOVAs 327 Understanding the Interaction Term 331 Plotting Results from ANCOVAs 333

Comparing Means 335 A Posteriori Comparisons 337 A Priori Contrasts 339

Model Selection Criteria 282 Model Selection Methods for Multiple Regression 283 Model Selection Methods in Path Analysis 284 Bayesian Model Selection 285

Bonferroni Corrections and the Problem of Multiple Tests 345 Summary 348

CHAPTER 11

The Analysis of Categorical Data 349

Summary 287

CHAPTER 10

Two-Way Contingency Tables 350

The Analysis of Variance 289 Symbols and Labels in ANOVA 290 ANOVA and Partitioning of the Sum of Squares 290 The Assumptions of ANOVA 295

Organizing the Data 350 Are the Variables Independent? 352 Testing the Hypothesis: Pearson’s Chi-square Test 354 An Alternative to Pearson’s Chi-Square: The G-Test 358

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Contents The Chi-square Test and the G-Test for R × C Tables 359 Which Test To Choose? 363

Multi-Way Contingency Tables 364 Organizing the Data 364 On to Multi-Way Tables! 368 Bayesian Approaches to Contingency Tables 375

Tests for Goodness-of-Fit 376 Goodness-of-Fit Tests for Discrete Distributions 376 Testing Goodness-of-Fit for Continuous Distributions: The Kolmogorov-Smirnov Test 380

Summary 382

CHAPTER 12

The Multivariate Normal Distribution 394 Testing for Multivariate Normality 396

Measurements of Multivariate Distance 398 Measuring Distances between Two Individuals 398 Measuring Distances between Two Groups 402 Other Measurements of Distance 402

Ordination 406 Principal Component Analysis 406 Factor Analysis 415 Principal Coordinates Analysis 418 Correspondence Analysis 421 Non-Metric Multidimensional Scaling 425 Advantages and Disadvantages of Ordination 427

Classification 429

The Analysis of Multivariate Data 383 Approaching Multivariate Data 383 The Need for Matrix Algebra 384

Comparing Multivariate Means 387 Comparing Multivariate Means of Two Samples: Hotelling’s T 2 Test 387 Comparing Multivariate Means of More Than Two Samples: A Simple MANOVA 390

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Cluster Analysis 429 Choosing a Clustering Method 430 Discriminant Analysis 433 Advantages and Disadvantages of Classification 437

Multivariate Multiple Regression 438 Redundancy Analysis 438

Summary 444

PART IV

Estimation CHAPTER 13

The Measurement of Biodiversity 449 Estimating Species Richness 450 Standardizing Diversity Comparisons through Random Subsampling 453

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Rarefaction Curves: Interpolating Species Richness 455 The Expectation of the Individual-Based Rarefaction Curve 459 Sample-Based Rarefaction Curves: Massachusetts Ants 461 Species Richness versus Species Density 465

The Statistical Comparison of Rarefaction Curves 466 Assumptions of Rarefaction 467

Asymptotic Estimators: Extrapolating Species Richness 470 Rarefaction Curves Redux: Extrapolation and Interpolation 476

Estimating Species Diversity and Evenness 476 Hill Numbers 479

Software for Estimation of Species Diversity 481 Summary 482

Occupancy of More than One Species 493 A Hierarchical Model for Parameter Estimation and Modeling 495 Occupancy Models for Open Populations 501 Dynamic Occupancy of the Adelgid in Massachusetts 505

Estimating Population Size 506 Mark-Recapture: The Basic Model 507 Mark-Recapture Models for Open Populations 516

Occupancy Modeling and Mark-Recapture: Yet More Models 518 Sampling for Occupancy and Abundance 519 Software for Estimating Occupancy and Abundance 521 Summary 522

APPENDIX

Matrix Algebra for Ecologists 523 CHAPTER 14

Detecting Populations and Estimating their Size 483

Glossary 535 Literature Cited 565 Index 583

Occupancy 485 The Basic Model: One Species, One Season, Two Samples at a Range of Sites 487

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Preface What is the “added value” for readers in purchasing this Second Edition? For this Second Edition, we have added a new Part 4, on “Estimation,” which includes two chapters: “The Measurement of Biodiversity” (Chapter 13), and “Detecting Populations and Estimating their Size” (Chapter 14). These two new chapters describe measurements and methods that are beginning to be widely used and that address questions that are central to the disciplines of community ecology and demography. Some of the methods we describe have been developed only in the last 10 years, and many of them continue to evolve rapidly. The title of this new section, “Estimation,” reflects an ongoing shift in ecology from testing hypotheses to estimating parameters. Although Chapter 3 describes summary measures of location and spread, most of the focus of the First Edition—and of many ecological publications—is on hypothesis testing (Chapter 4). In the two new chapters, we are more concerned with the process of estimation itself, although the resulting estimators of biodiversity and population size certainly can be used in conventional statistical tests once they are calculated. It is useful to think of biodiversity and population sizes as latent or unknown “state variables.” We want to measure these things, but it is impossible to find and tag all of the trout in a stream or discover all of the species of ants that lurk in a bog. Instead, we work only with small samples of diversity, or limited collections of marked animals, and we use these samples to estimate the “true” value of the underlying state variables. The two new chapters are united not only by their focus on estimation, but also by their common underlying statistical framework: methods for asymptotic estimators of species richness discussed in Chapter 13 were derived from methods developed for mark-recapture studies discussed in Chapter 14. We hope the methods described in these two chapters will provide some fresh statistical tools for ecologists to complement the more classical topics we cover in Chapters 1–12. Chapter 13 begins with what seems like a relatively simple question: how many species are there in an assemblage? The central problem is that nature is

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(still) vast, our sample of its diversity is tiny, and there are always many rare and undetected species. Moreover, because the number of species we count is heavily influenced by the number of individuals and samples we have collected, the data must be standardized for meaningful comparisons. We describe methods for both interpolation (rarefaction) and extrapolation (asymptotic species richness estimators) to effectively standardize biodiversity comparisons. We also advocate the use of Hill numbers as a class of diversity indices that have useful statistical properties, although they too are subject to sampling effects. We thank Anne Chao, Rob Colwell, and Lou Jost for their ongoing collaboration, as well as their insights, extended correspondence, and important contributions to the literature on biodiversity estimation. In Chapter 14, we delve deeper into the problem of incomplete detection, addressing two main questions. First, how do we estimate the probability that we can detect a species, given that it is actually present at a site? Second, for a species that is present at a site, how large is its population? Because we have a limited sample of individuals that we can see and count, we must first estimate the detection probability of an individual species before we can estimate the size of its population. Estimates of occupancy probability and population size are used in quantitative models for the management of populations of both rare and exploited species. And, just as we use species richness estimators to extrapolate beyond our sample, we use estimates from occupancy models and markrecapture studies to extrapolate and forecast what will happen to these populations as, for example, habitats are fragmented, fishing pressure increases, or the climate changes. We thank Elizabeth Crone, Dave Orwig, Evan Preisser, and Rui Zhang for letting us use some of their unpublished data in this chapter and for discussing their analyses with us. Bob Dorazio and Andy Royle discussed these models with us; they, along with Evan Cooch, have developed state-of-the-art open-source software for occupancy modeling and mark-recapture analysis. In this edition, we have also corrected numerous minor errors that astute readers have pointed out to us over the past 8 years. We are especially grateful for the careful reading of Victor Lemes Landeiro, who, together with his colleagues Fabricio Beggiato Baccaro, Helder Mateus Viana Espirito Santo, Miriam Plaza Pinto, and Murilo Sversut Dias, translated the entire book into Portuguese. Specific contributions of these individuals, and all the others who sent us comments and pointed out errors, are noted in the Errata section of the book’s website (harvardforest.fas.harvard.edu/ellison/publications/primer/errata). Although we have corrected all errors identified in the first edition, others undoubtedly remain (another problem in detection probability); please contact us if you find any. All code posted on the Data and Code section of the book’s website (harvardforest.fas.harvard.edu/ellison/publications/primer/datafiles) that we used

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for the analyses in the book has been updated and reworked in the R programming language (r-project.org). In the time since we wrote the first edition, R has become the de facto standard software for statistical analysis and is used by most ecologists and statisticians for their day-to-day analytical work. We have provided some R scripts for our figures and analyses, but they have minimal annotation, and there are already plenty of excellent resources available for ecological modeling and graphics with R. Last but not least, we thank Andy Sinauer, Azelie Aquadro, Joan Gemme, Chris Small, Randy Burgess, and the entire staff at Sinauer Associates for carefully editing our manuscript pages and turning them into an attractive book.1

Preface to the First Edition Why another book on statistics? The field is already crowded with texts on biometry, probability theory, experimental design, and analysis of variance. The short answer is that we have yet to find a single text that meets the two specific needs of ecologists and environmental scientists. The first need is a general introduction to probability theory including the assumptions of statistical inference and hypothesis testing. The second need is a detailed discussion of specific designs and analyses that are typically encountered in ecology. This book reflects several collective decades of teaching statistics to undergraduate and graduate students at the University of Oklahoma, the University of Vermont, Mount Holyoke College, and the Harvard Forest. The book represents our personal perspective on what is important at the interface of ecology and statistics and the most effective way to teach it. It is the book we both wish we had in our hands when we started studying ecology in college. A “primer” suggests a short text, with simple messages and safe recommendations for users. If only statistics were so simple! The subject is vast, and new methods and software tools for ecologists appear almost monthly. In spite of the book’s length (which stunned even us), we still view it as a distillation of statistical knowledge that is important for ecologists. We have included both classic expositions as well as discussions of more recent methods and techniques. We hope there will be material that will appeal to both the neophyte and the experienced researcher. As in art and music, there is a strong element of style and personal preference in the choice and application of statistics. We have explained our own preferences for certain kinds of analyses and designs, but readers and users of statistics will ultimately have to choose their own set of statistical tools.

And for dealing with all of the footnotes!

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How To Use This Book

In contrast to many statistics books aimed at biologists or ecologists—“biometry” texts—this is not a book of “recipes,” nor is it a set of exercises or problems tied to a particular software package that is probably already out of date. Although the book contains equations and derivations, it is not a formal statistical text with detailed mathematical proofs. Instead, it is an exposition of basic and advanced topics in statistics that are important specifically for ecologists and environmental scientists. It is a book that is meant to be read and used, perhaps as a supplement to a more traditional text, or as a stand-alone text for students who have had at least a minimal introduction to statistics. We hope this book will also find a place on the shelf (or floor) of environmental professionals who need to use and interpret statistics daily but who may not have had formal training in statistics or have ready access to helpful statisticians. Throughout the book we make extensive use of footnotes. Footnotes give us a chance to greatly expand the material and to talk about more complex issues that would bog down the flow of the main text. Some footnotes are purely historical, others cover mathematical and statistical proofs or details, and others are brief essays on topics in the ecological literature. As undergraduates, we both were independently enamored of Hutchinson’s classic An Introduction to Population Biology (Yale University Press 1977). Hutchinson’s liberal use of footnotes and his frequent forays into history and philosophy served as our stylistic model. We have tried to strike a balance between being concise and being thorough. Many topics recur in different chapters, although sometimes in slightly different contexts. Figure and table legends are somewhat expansive because they are meant to be readable without reference to the text. Because Chapter 12 requires matrix algebra, we provide a brief Appendix that covers the essentials of matrix notation and manipulation. Finally, we have included a comprehensive glossary of terms used in statistics and probability theory. This glossary may be useful not only for reading this book, but also for deciphering statistical methods presented in journal articles. Mathematical Content

Although we have not shied away from equations when they are needed, there is considerably less math here than in many intermediate level texts. We also try to illustrate all methods with empirical analyses. In almost all cases, we have used data from our own studies so that we can illustrate the progression from raw data to statistical output. Although the chapters are meant to stand alone, there is considerable crossreferencing. Chapters 9–12 cover the traditional topics of many biometry texts and contain the heaviest concentration of formulas and equations. Chapter 12,

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on multivariate methods, is the most advanced in the book. Although we have tried to explain multivariate methods in plain English (no easy task!), there is no way to sugar-coat the heavy dose of matrix algebra and new vocabulary necessary for multivariate analysis. Coverage and Topics

Statistics is an evolving field, and new methods are always being developed to answer ecological questions. This text, however, covers core material that we believe is foundational for any statistical analysis. The book is organized around design-based statistics—statistics that are to be used with designed observational and experimental studies. An alternative framework is model-based statistics, including model selection criteria, likelihood analysis, inverse modeling, and other such methods. What characterizes the difference between these two approaches? In brief, design-based analyses primarily address the problem of P(data | model): assessing the probability of the data given a model that we have specified. An important element of design-based analysis is being explicit about the underlying model. In contrast, model-based statistics address the problem of P (model | data)—that is, assessing the probability of a model given the data that are present. Model-based methods are often most appropriate for large datasets that may not have been collected according to an explicit sampling strategy. We also discuss Bayesian methods, which somewhat straddle the line between these two approaches. Bayesian methods address P(model | data), but can be used with data from designed experiments and observational studies. The book is divided into three parts. Part I discusses the fundamentals of probability and statistical thinking. It introduces the logic and language of probability (Chapter 1), explains common statistical distributions used in ecology (Chapter 2), and introduces important measures of location and spread (Chapter 3). Chapter 4 is more of a philosophical discussion about hypothesis testing, with careful explanations of Type I and Type II errors and of the ubiquitous statement that “P < 0.05.” Chapter 5 closes this section by introducing the three major paradigms of statistical analysis (frequentist, Monte Carlo, and Bayesian), illustrating their use through the analysis of a single dataset. Part II discusses how to successfully design and execute observational studies and field experiments. These chapters contain advice and information that is to be used before any data are collected in the field. Chapter 6 addresses the practical problems of articulating the reason for the study, setting the sample size, and dealing with independence, replication, randomization, impact studies, and environmental heterogeneity. Chapter 7 is a bestiary of experimental designs. However, in contrast to almost all other texts, there are almost no equations in this

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chapter. We discuss the strengths and weaknesses of different designs without getting bogged down in equations or analysis of variance (ANOVA) tables, which have mostly been quarantined in Part III. Chapter 8 addresses the problem of how to curate and manage the data once it is collected. Transformations are introduced in this chapter as a way to help screen for outliers and errors in data transcription. We also stress the importance of creating metadata—documentation that accompanies the raw data into long-term archival storage. Part III discusses specific analyses and covers the material that is the main core of most statistics texts. These chapters carefully develop many basic models, but also try to introduce some more current statistical tools that ecologists and environmental scientists find useful. Chapter 9 develops the basic linear regression model and also introduces more advanced topics such as non-linear, quantile, and robust regression, and path analysis. Chapter 10 discusses ANOVA (analysis of variance) and ANCOVA (analysis of covariance) models, highlighting which ANOVA model is appropriate for particular sampling or experimental designs. This chapter also explains how to use a priori contrasts and emphasizes plotting ANOVA results in a meaningful way to understand main effects and interaction terms. Chapter 11 discusses categorical data analysis, with an emphasis on chisquare analysis of one- and two-way contingency tables and log-linear modeling of multi-way contingency tables. The chapter also introduces goodness-of-fit tests for discrete and continuous variables. Chapter 12 introduces a variety of multivariate methods for both ordination and classification. The chapter closes with an introduction to redundancy analysis, a multivariate analog of multiple regression. About the Cover

The cover is an original still life by the ecologist Elizabeth Farnsworth, inspired by seventeenth-century Dutch still-life paintings. Netherlands painters, including De Heem, Claesz, and Kalf, displayed their skill by arranging and painting a wide range of natural and manmade objects, often with rich allusions and hidden symbolism. Elizabeth’s beautiful and biologically accurate renderings of plants and animals follow a long tradition of biological illustrators such as Dürer and Escher. In Elizabeth’s drawing, the centerpiece is a scrolled pen-and-paper illustration of Bayes’ Theorem, an eighteenth-century mathematical exposition that lies at the core of probability theory. Although modern Bayesian methods are computationally intensive, an understanding of Bayes’ Theorem still requires pen-and-paper contemplation. Dutch still lifes often included musical instruments, perhaps symbolizing the Pythagorean beauty of music and numbers. Elizabeth’s drawing incorporates Aaron’s bouzouki, as the writing of this book was interwoven with many sessions of acoustic music with Aaron and Elizabeth.

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The human skull is the traditional symbol of mortality in still life paintings. Elizabeth has substituted a small brain coral, in honor of Nick’s early dissertation work on the ecology of subtropical gorgonians. In light of the current worldwide collapse of coral reefs, the brain coral seems an appropriate symbol of mortality. In the foreground, an Atta ant carries a scrap of text with a summation sign, reminding us of the collective power of individual ant workers whose summed activities benefit the entire colony. As in early medieval paintings, the ant is drawn disproportionately larger than the other objects, reflecting its dominance and importance in terrestrial ecosystems. In the background is a specimen of Sarracenia minor, a carnivorous plant from the southeastern U.S. that we grow in the greenhouse. Although Sarracenia minor is too rare for experimental manipulations in the field, our ongoing studies of the more common Sarracenia purpurea in the northeastern U.S. may help us to develop effective conservation strategies for all of the species in this genus. The final element in this drawing is a traditional Italian Renaissance chicken carafe. In the summer of 2001, we were surveying pitcher plants in remote bogs of the Adirondack Mountains. At the end of a long field day, we found ourselves in a small Italian restaurant in southern Vermont. Somewhat intemperately, we ordered and consumed an entire liter-sized chicken carafe of Chianti. By the end of the evening, we had committed ourselves to writing this book, and “The Chicken”—as we have always called it—was officially hatched.

Acknowledgments We thank several colleagues for their detailed suggestions on different chapters: Marti Anderson (11, 12), Jim Brunt (8), George Cobb (9, 10), Elizabeth Farnsworth (1–5), Brian Inouye (6, 7), Pedro Peres-Neto (11,12), Catherine Potvin (9, 10), Robert Rockwell (1–5), Derek Roff (original book proposal), David Skelly (original book proposal), and Steve Tilley (1–5). Laboratory discussion groups at the Harvard Forest and University of Vermont also gave us feedback on many parts of the book. Special kudos go to Henry Horn, who read and thoroughly critiqued the entire manuscript. We thank Andy Sinauer, Chris Small, Carol Wigg, Bobbie Lewis, and Susan McGlew of Sinauer Associates, and, especially, Michele Ruschhaupt of The Format Group for transforming pages of crude word-processed text into an elegant and pleasing book. The National Science Foundation supports our research on pitcher plants, ants, null models, and mangroves, all of which appear in many of the examples in this book. The University of Vermont and the Harvard Forest, our respective home institutions, provide supportive environments for book writing.

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PART I

Fundamentals of Probability and Statistical Thinking

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CHAPTER 1

An Introduction to Probability

In this chapter, we develop basic concepts and definitions required to understand probability and sampling. The details of probability calculations lie behind the computation of all statistical significance tests. Developing an appreciation for probability will help you design better experiments and interpret your results more clearly. The concepts in this chapter lay the foundations for the use and understanding of statistics. They are far more important, actually, than some of the detailed topics that will follow. This chapter provides the necessary background to address the critical questions such as “What does it mean when you read in a scientific paper that the means of two samples were significantly different at P = 0.003?” or “What is the difference between a Type I and a Type II statistical error?” You would be surprised how often scientists with considerable experience in statistics are unable to clearly answer these questions. If you can understand the material in this introductory chapter, you will have a strong basis for your study of statistics, and you will always be able to correctly interpret statistical material in the scientific literature—even if you are not familiar with the details of the particular test being used. In this chapter, we also introduce you to the problem of measurement and quantification, processes that are essential to all sciences. We cannot begin to investigate phenomena scientifically unless we can quantify processes and agree on a common language with which to interpret our measurements. Of course, the mere act of quantification does not by itself make something a science: astrology and stock market forecasting use a lot of numbers, but they don’t qualify as sciences. One conceptual challenge for students of ecology is translating their “love of nature” into a “love of pattern.” For example, how do we quantify patterns of plant and animal abundance? When we take a walk in the woods, we find ourselves asking a stream of questions such as: “What is our best estimate for the density of Myrmica (a common forest ant) colonies in this forest? Is it 1 colony/m2? 10 colonies/m2? What would be the best way to measure Myrmica density? How does Myrmica density vary in different parts of the woods? What mechanisms or

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CHAPTER 1 An Introduction to Probability

hypotheses might account for such variation?” And, finally, “What experiments and observations could we make to try and test or falsify these hypotheses?” But once we have “quantified nature,” we still have to summarize, synthesize, and interpret the data we have collected. Statistics is the common language used in all sciences to interpret our measurements and to test and discriminate among our hypotheses (e.g., Ellison and Dennis 2010). Probability is the foundation of statistics, and therefore is the starting point of this book.

What Is Probability? If the weather forecast calls for a 70% chance of rain, we all have an intuitive idea of what that means. Such a statement quantifies the probability, or the likely outcome, of an event that we are unsure of. Uncertainty exists because there is variation in the world, and that variation is not always predictable. Variation in biological systems is especially important; it is impossible to understand basic concepts in ecology, evolution, and environmental science without an appreciation of natural variation.1 Although we all have a general understanding of probability, defining it precisely is a different matter. The problem of defining probability comes into sharper focus when we actually try to measure probabilities of real events.

Measuring Probability The Probability of a Single Event: Prey Capture by Carnivorous Plants

Carnivorous plants are a good system for thinking about the definition of probability. The northern pitcher plant Sarracenia purpurea captures insect prey in leaves that fill with rainwater.2 Some of the insects that visit a pitcher plant fall into 1

Variation in traits among individuals is one of the key elements of the theory of evolution by natural selection. One of Charles Darwin’s (1809–1882) great intellectual contributions was to emphasize the significance of such variation and to break free from the conceptual straightjacket of the typological “species concept,” which treated species as fixed, static entities with well-defined, unchanging boundaries.

Charles Darwin 2

Carnivorous plants are some of our favorite study organisms. They have fascinated biologists since Darwin first proved that they can absorb nutrients from insects in 1875. Carnivorous plants have many attributes that make them model ecological systems (Ellison and Gotelli 2001; Ellison et al. 2003).

Pitcher plant (Sarracenia purpurea)

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Measuring Probability

the pitcher and drown; the plant extracts nutrients from the decomposing prey. Although the trap is a marvelous evolutionary adaptation for life as a carnivore, it is not terribly efficient, and most of the insects that visit a pitcher plant escape. How could we estimate the probability that an insect visit will result in a successful capture? The most straightforward way would be to keep careful track of the number of insects that visit a plant and the number that are captured. Visiting a plant is an example of what statisticians call an event. An event is a simple process with a well-recognized beginning and end. In this simple universe, visiting a plant is an event that can result in two outcomes: escape or capture. Prey capture is an example of a discrete outcome because it can be assigned a positive integer. For example, you could assign a 1 for a capture and a 2 for an escape. The set formed from all the possible outcomes of an event is called a sample space.3 Sample spaces or sets made up of discrete outcomes are called discrete sets because their outcomes are all countable. 4 3

Even in this simple example, the definitions are slippery and won’t cover all possibilities. For example, some flying insects may hover above the plant but never touch it, and some crawling insects may explore the outer surface of the plant but not cross the lip and enter the pitcher. Depending on how precise we care to be, these possibilities may or may not be included in the sample space of observations we use to determine the probability of being captured. The sample space establishes the domain of inference from which we can draw conclusions. 4

The term countable has a very specific meaning in mathematics. A set is considered countable if every element (or outcome) of that set can be assigned a positive integer value. For example, we can assign to the elements of the set Visit the integers 1 (= capture) and 2 (= escape). The set of integers itself is countable, even though there are infinitely many integers. In the late 1800s, the mathematician Georg Ferdinand Ludwig Philipp Cantor G. F. L. P. Cantor (1845–1918), one of the founders of what has come to be called set theory, developed the notion of cardinality to describe such countability. Two sets are considered to have the same cardinality if they can be put in one-to-one correspondence with each other. For example, the elements of the set of all even integers {2, 4, 6, …} can each be assigned an element of the set of all integers (and vice versa). Sets that have the same cardinality as the integers are said to have cardinality 0, denoted as ℵ0. Cantor went on to prove that the set of rational numbers (numbers that can be written as the quotient of any two integers) also has cardinality ℵ0, but the set of irrational numbers (numbers that cannot be written as the quotient of any two integers, such as 冑苳 2 ) is not countable. The set of irrational numbers has cardinality 1, and is denoted ℵ1. One of Cantor’s most famous results is that there is a one-to-one correspondence between the set of all points in the small interval [0,1] and the set of all points in n-dimensional space (both have cardinality 1). This result, published in 1877, led him to write to the mathematician Julius Wilhelm Richard Dedekind, “I see it, but I don’t believe it!” Curiously, there are types of sets that have higher cardinality than 1, and in fact there are infinitely many cardinalities. Still weirder, the set of cardinal numbers {ℵ0, ℵ1, ℵ2,…} is itself a countably infinite set (its cardinality = 0)!

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CHAPTER 1 An Introduction to Probability

Each insect that visits a plant is counted as a trial. Statisticians usually refer to each trial as an individual replicate, and refer to a set of trials as an experiment.5 We define the probability of an outcome as the number of times that outcome occurs divided by the number of trials. If we watched a single plant and recorded 30 prey captures out of 3000 visits, we would calculate the probability as number of captures number of visits or 30 in 3000, which we can write as 30/3000, or 0.01. In more general terms, we calculate the probability P that an outcome occurs to be P=

number of outcomes number of trials

(1.1)

By definition, there can never be more outcomes than there are trials, so the numerator of this fraction is never larger than the denominator, and therefore 0.0 ≤ P ≤ 1.0 In other words, probabilities are always bounded by a minimum of 0.0 and a maximum of 1.0. A probability of 0.0 represents an event that will never happen, and a probability of 1.0 represents an event that will always happen. However, even this estimate of probability is problematic. Most people would say that the probability that the sun will rise tomorrow is a sure thing (P = 1.0). Certainly if we assiduously recorded the rising of the sun each morning, we would find that it always does, and our measurements would, in fact, yield a probability estimate of 1.0. But our sun is an ordinary star, a nuclear reactor that releases energy when hydrogen atoms fuse to form helium. After about 10 billion years, all the hydrogen fuel is used up, and the star dies. Our sun is a middle-aged star, so if you could keep observing for another 5 billion years, one morning the sun would no longer rise. And if you started your observations at that point, your estimate of the probability of a sunrise would be 0.0. So, is the probability that the sun rises each day really 1.0? Something less than 1.0? 1.0 for now, but 0.0 in 5 billion years? 5

This statistical definition of an experiment is less restrictive than the conventional definition, which describes a set of manipulated subjects (the treatment) and an appropriate comparison group (the control). However, many ecologists use the term natural experiment to refer to comparisons of replicates that have not been manipulated by the investigator, but that differ naturally in the quantity of interest (e.g., islands with and without predators; see Chapter 6).

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Measuring Probability

This example illustrates that any estimate or measure of probability is completely contingent on how we define the sample space—the set of all possible events that we use for comparison. In general, we all have intuitive estimates or guesses for probabilities for all kinds of events in daily life. However, to quantify those guesses, we have to decide on a sample space, take samples, and count the frequency with which certain events occur. Estimating Probabilities by Sampling

In our first experiment, we watched 3000 insects visit a plant; 30 of them were captured, and we calculated the probability of capture as 0.01. Is this number reasonable, or was our experiment conducted on a particularly good day for the plants (or a bad day for the insects)? There will always be variability in these sorts of numbers. Some days, no prey are captured; other days, there are several captures. We could determine the true probability of capture precisely if we could watch every insect visiting a plant at all times of day or night, every day of the year. Each time there was a visit, we would determine whether the outcome Capture occurred and accordingly update our estimate of the probability of capture. But life is too short for this. How can we estimate the probability of capture without constantly monitoring plants? We can efficiently estimate the probability of an event by taking a sample of the population of interest. For example, every week for an entire year we could watch 1000 insects visit plants on a single day. The result is a set of 52 samples of 1000 trials each, and the number of insects captured in each sample. The first three rows of this 52-row dataset are shown in Table 1.1. The probabilities of capture differ on the different days, but not by very much; it appears to be relatively uncommon for insects to be captured while visiting pitcher plants.

TABLE 1.1 Sample data for number of insect captures per 1000 visits to the carnivorous plant Sarracenia purpurea ID number

1 2 3 ⯗ 52

Observation date

Number of insect captures

June 1, 1998 June 8, 1998 June 15, 1998 ⯗ May 24, 1999

10 13 12 ⯗ 11

In this hypothetical dataset, each row represents a different week in which a sample was collected. If the sampling was conducted once a week for an entire year, the dataset would have exactly 52 rows. The first column indicates the ID number, a unique consecutive integer (1 to 52) assigned to each row of the dataset. The second column gives the sample date, and the third column gives the number of insect captures out of 1000 insect visits that were observed. From this dataset, the probability of capture for any single sample can be estimated as the number of insect captures/1000. The data from all 52 capture numbers can be plotted and summarized as a histogram that illustrates the variability among the samples (see Figure 1.1).

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CHAPTER 1 An Introduction to Probability

Figure 1.1 A histogram of capture frequencies. Each week for an entire year, an investigator might observe 1000 insect visits to a carnivorous pitcher plant and count how many times out of those visits the plant captures an insect (see Table 1.1). The collected data from all 52 observations (one per week) can be plotted in the form of a histogram. The number of captures observed ranges from as few as 4 one week (bar at left end) to as many as 20 another week (bar at right end); the average of all 52 observations is 10.3 captures per 1000 visits.

Average frequency of capture = 10.3 per 1000 visits

8 Number of observations

0 0

6 8 10 12 14 16 18 Number of insects captured per 1000 visits

In Figure 1.1, we concisely summarize the results of one year’s worth of samples6 in a graph called a histogram. In this particular histogram, the numbers on the horizontal axis, or x-axis, indicate how many insects were captured while visiting plants. The numbers range from 4 to 20, because in the 52 samples, there were days when only 4 insects were captured, days when 20 insects were captured, and some days in between. The numbers on the vertical axis, or y-axis indicate the frequency: the number of trials that resulted in a particular outcome. For example, there was only one day in which there were 4 captures recorded, two days in which there were 6 captures recorded, and five days in which 9 captures were recorded. One way we can estimate the probability of capture is to take the average of all the samples. In other words, we would calculate the average number of captures in each of our 52 samples of 1000 visits. In this example, the average number of captures out of 1000 visits is 10.3. Thus, the probability of being captured is 10.3/1000 = 0.0103, or just over one in a hun6

Although we didn’t actually conduct this experiment, these sorts of data were collected by Newell and Nastase (1998), who videotaped insect visits to pitcher plants and recorded 27 captures out of 3308 visits, with 74 visits having an unknown fate. For the analyses in this chapter, we simulated data using a random-number generator in a computer spreadsheet. For this simulation, we set the “true” probability of capture at 0.01, the sample size at 1000 visits, and the number of samples at 52. We then used the resulting data to generate the histogram shown in Figure 1.1. If you conduct the same simulation experiment on your computer, you should get similar results, but there will be some variability depending on how your computer generates random data. We discuss simulation experiments in more detail in Chapters 3 and 5.

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Problems in the Definition of Probability

dred. We call this average the expected value of the probability, or expectation, and denote it as E(P). Distributions like the one shown in Figure 1.1 are often used to describe the results of experiments in probability. We return to them in greater detail in Chapter 3.

Problems in the Definition of Probability Most statistics textbooks very briefly define probability just as we have done: the (expected) frequency with which events occur. The standard textbook examples are that the toss of a fair coin has a 0.50 probability (equals a 50% chance) of landing heads,7 or that each face of a six-sided fair die has a 1 in 6 chance of turning up. But let us consider these examples more carefully. Why do we accept these values as the correct probabilities? We accept these answers because of our view of how coins are tossed and dice are rolled. For example, if we were to balance the coin on a tabletop with “heads” facing toward us, and then tip the coin gently forward, we could all agree that the probability of obtaining heads would no longer be 0.50. Instead, the probability of 0.50 only applies to a coin that is flipped vigorously into the air.8 Even here, however, we should recognize that the 0.50 probability really represents an estimate based on a minimal amount of data. For example, suppose we had a vast array of high-tech microsensors attached to the surface of the coin, the muscles of our hand, and the walls of the room we are in. These sensors detect and quantify the exact amount of torque in our toss, the temperature and air turbulence in the room, the micro-irregularities on the surface of the coin, and the microeddies of air turbulence that are generated as the coin flips. If all these data were instantaneously streamed into a fast 7

In a curious twist of fate, one of the Euro coins may not be fair. The Euro coin, introduced in 2002 by 12 European states, has a map of Europe on the “tails” side, but each country has its own design on the “heads” side. Polish statisticians Tomasz Gliszcynski and Waclaw Zawadowski flipped (actually, spun) the Belgian Euro 250 times, and it came up heads 56% of the time (140 heads). They attributed this result to a heavier embossed image on the “heads” side, but Howard Grubb, a statistician at the University of Reading, points out that for 250 trials, 56% is “not significantly different” from 50%. Who’s right? (as reported by New Scientist: www.newscientist.com/, January 2, 2002). We return to this example in Chapter 11.

Gamblers and cardsharps try to thwart this model by using loaded coins or dice, or by carefully tossing coins and dice in a way that favors one of the faces but gives the appearance of randomness. Casinos work vigilantly to ensure that the random model is enforced; dice must be shaken vigorously, roulette wheels are spun very rapidly, and blackjack decks are frequently reshuffled. It isn’t necessary for the casino to cheat. The payoff schedules are calculated to yield a handsome profit, and all that the casino must do is to ensure that customers have no way to influence or predict the outcome of each game.

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CHAPTER 1 An Introduction to Probability

computer, we could develop a very complex model that describes the trajectory of the coin. With such information, we could predict, much more accurately, which face of the coin would turn up. In fact, if we had an infinite amount of data available, perhaps there would be no uncertainty in the outcome of the coin toss.9 The trials that we use to estimate the probability of an event may not be so similar after all. Ultimately, each trial represents a completely unique set of conditions; a particular chain of cause and effect that determines entirely whether the coin will turn up heads or tails. If we could perfectly duplicate that set of conditions; there would be no uncertainty in the outcome of the event, and no need to use probability or statistics at all!10 9

This last statement is highly debatable. It assumes that we have measured all the right variables, and that the interactions between those variables are simple enough that we can map out all of the contingencies or express them in a mathematical relationship. Most scientists believe that complex models will be more accurate than simple ones, but that it may not be possible to eliminate all uncertainty (Lavine 2010). Note also that complex models don’t help us if we cannot measure the variables they include; the Werner Heisenberg technology needed to monitor our coin tosses isn’t likely to exist anytime soon. Finally, there is the more subtle problem that the very act of measuring things in nature may alter the processes we are trying to study. For example, suppose we want to quantify the relative abundance of fish species near the deep ocean floor, where sunlight never penetrates. We can use underwater cameras to photograph fish and then count the number of fish in each photograph. But if the lights of the camera attract some fish species and repel others, what, exactly, have we measured? The Erwin Schrödinger Heisenberg Uncertainty Principle (named after the German physicist Werner Heisenberg, 1901–1976) in physics says that it is impossible to simultaneously measure the position and momentum of an electron: the more accurately you measure one of those quantities, the less accurately you can measure the other. If the very act of observing fundamentally changes processes in nature, then the chain of cause and effect is broken (or at least badly warped). This concept was elaborated by Erwin Schrödinger (1887–1961), who placed a (theoretical) cat and a vial of cyanide in a quantum box. In the box, the cat can be both alive and dead simultaneously, but once the box is opened and the cat observed, it is either alive or eternally dead. 10

Many scientists would embrace this lack of uncertainty. Some of our molecular biologist colleagues would be quite happy in a world in which statistics are unnecessary. When there is uncertainty in the outcome of their experiments, they often assume that it is due to bad experimental technique, and that eliminating measurement error and contamination will lead to clean and repeatable data that are correct. Ecologists and field biologists usually are more sanguine about variation in their systems. This is not necessarily because ecological systems are inherently more noisy than molecular ones; rather, ecological data may be more variable than molecular data because they are often collected at different spatial and temporal scales.

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The Mathematics of Probability

Returning to the pitcher plants, it is obvious that our estimates of capture probabilities will depend very much on the details of the trials. The chances of capture will differ between large and small plants, between ant prey and fly prey, and between plants in sun and plants in shade. And, in each of these groups, we could make further subdivisions based on still finer details about the conditions of the visit. Our estimates of probabilities will depend on how broadly or how narrowly we limit the kinds of trials we will consider. To summarize, when we say that events are random, stochastic, probabilistic, or due to chance, what we really mean is that their outcomes are determined in part by a complex set of processes that we are unable or unwilling to measure and will instead treat as random. The strength of other processes that we can measure, manipulate, and model represent deterministic or mechanistic forces. We can think of patterns in our data as being determined by mixtures of such deterministic and random forces. As observers, we impose the distinction between deterministic and random. It reflects our implicit conceptual model about the forces in operation, and the availability of data and measurements that are relevant to these forces.11

The Mathematics of Probability In this section, we present a brief mathematical treatment of how to calculate probabilities. Although the material in this section may seem tangential to your quest (Are the data “significant”?), the correct interpretation of statistical results hinges on these operations. Defining the Sample Space

As we illustrated in the example of prey capture by carnivorous plants, the probability P of being captured (the outcome) while visiting a plant (the event) is 11

If we reject the notion of any randomness in nature, we quickly move into the philosophic realm of mysticism: “What, then, is this Law of Karma? The Law, without exception, which rules the whole universe, from the invisible, imponderable atom to the suns; … and this law is that every cause produces its effect, without any possibility of delaying or annulling that effect, once the cause begins to operate. The law of causation is everywhere supreme. This is the Law of Karma; Karma is the inevitable link between cause and effect” (Arnould 1895). Western science doesn’t progress well by embracing this sort of all-encompassing complexity. It is true that many scientific explanations for natural phenomena are complicated. However, those explanations are reached by first eschewing all that complexity and posing a null hypothesis. A null hypothesis tries to account for patterns in the data in the simplest way possible, which often means initially attributing variation in the data to randomness (or measurement error). If that simple null hypothesis can be rejected, we can move on to entertain more complex hypotheses. See Chapter 4 for more details on null hypotheses and hypothesis testing, and Beltrami (1999) for a discussion of randomness.

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CHAPTER 1 An Introduction to Probability

defined simply as the number of captures divided by the number of visits (the number of trials) (Equation 1.1). Let’s examine in detail each of these terms: outcome, event, trial, and probability. First, we need to define our universe of possible events, or the sample space of interest. In our first example, an insect could successfully visit a plant and leave, or it could be captured. These two possible outcomes form the sample space (or set), which we will call Visit: Visit = {(Capture), (Escape)} We use curly braces {} to denote the set and parentheses () to denote the events of a set. The objects within the parentheses are the outcome(s) of the event. Because there are only two possible outcomes to a visit, if the probability of the outcome Capture is 1 in 1000, or 0.001, then the probability of Escape is 999 in 1000, or 0.999. This simple example can be generalized to the First Axiom of Probability: Axiom 1: The sum of all the probabilities of outcomes within

a single sample space = 1.0. We can write this axiom as n

∑ P(Ai ) = 1.0 i=1

which we read as “the sum of the probabilities of all outcomes Ai equals 1.0.” The “sideways W” (actually the capital Greek letter sigma) is a summation sign, the shorthand notation for adding up the elements that follow. The subscript i indicates the particular element and says that we are to sum up the elements from i = 1 to n, where n is the total number of elements in the set. In a properly defined sample space, we say that the outcomes are mutually exclusive (an individual is either captured or it escapes), and that the outcomes are exhaustive (Capture or Escape are the only possible outcomes of the event). If the events are not mutually exclusive and exhaustive, the probabilities of events in the sample space will not sum to 1.0. In many cases, there will be more than just two possible outcomes of an event. For example, in a study of reproduction by the imaginary orange-spotted whirligig beetle, we found that each of these beasts always produces exactly 2 litters, with between 2 and 4 offspring per litter. The lifetime reproductive success of an orange-spotted whirligig can be described as an outcome (a,b), where a represents the number of offspring in the first litter and b the number of offspring in the second litter. The sample space Fitness consists of all the possible lifetime reproductive outcomes that an individual could achieve: Fitness = {(2,2), (2,3), (2,4), (3,2), (3,3), (3,4), (4,2), (4,3), (4,4)}

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The Mathematics of Probability

Because each whirligig can give birth to only 2, 3, or 4 offspring in a litter, these 9 pairs of integers are the only possible outcomes. In the absence of any other information, we initially make the simplifying assumption that the probabilities of each of these different reproductive outcomes are equal. We use the definition of probability from Equation 1.1 (number of outcomes/number of trials) to determine this value. Because there are 9 possible outcomes in this set, P(2,2) = P(2,3) = … = P(4,4) = 1/9. Notice also that these probabilities obey Axiom 1: the sum of the probabilities of all outcomes = 1/9 + 1/9 + 1/9 + 1/9 + 1/9 + 1/9 + 1/9 + 1/9 + 1/9 = 1.0. Complex and Shared Events: Combining Simple Probabilities

Once probabilities of simple events are known or have been estimated, we can use them to measure probabilities of more complicated events. Complex events are composites of simple events in the sample space. Shared events are multiple simultaneous occurrences of simple events in the sample space. The probabilities of complex and shared events can be decomposed into the sums or products of probabilities of simple events. However, it can be difficult to decide when probabilities are to be added and when they are to be multiplied. The answer can be found by determining whether the new event can be achieved by one of several different pathways (a complex event), or whether it requires the simultaneous occurrence of two or more simple events (a shared event). If the new event can occur via different pathways, it is a complex event and can be represented as an or statement: Event A or Event B or Event C. Complex events thus are said to represent the union of simple events. Probabilities of complex events are determined by summing the probabilities of simple events. In contrast, if the new event requires the simultaneous occurrence of several simple events, it is a shared event and can be represented as an and statement: Event A and Event B and Event C. Shared events therefore are said to represent the intersection of simple events. Probabilities of shared events are determined by multiplying probabilities together. COMPLEX EVENTS: SUMMING PROBABILITIES

The whirligig example can be used to illustrate the calculation of probabilities of complex events. Suppose we wish to measure the lifetime reproductive output of a whirligig beetle. We would count the total number of offspring that a whirligig produces in its lifetime. This number is the sum of the offspring of the two litters, which results in an integer number between 4 and 8, inclusive. How would we determine the probability that an orange-spotted whirligig produces 6 offspring? First, note that the event produces 6 offspring can occur in three ways: 6 offspring = {(2,4), (3,3), (4,2)}

and this complex event itself is a set.

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CHAPTER 1 An Introduction to Probability

Figure 1.2 Venn diagram illustrating the concept of sets. Each pair of numbers represents the number of offspring produced in two consecutive litters by an imaginary species of whirligig beetle. We assume that this beetle produces exactly 2, 3, or 4 offspring each time it reproduces, so these are the only integers represented in the diagram. A set in a Venn diagram is a ring that encompasses certain elements. The set Fitness contains all of the possible reproductive outcomes for two consecutive litters. Within the set Fitness is a smaller set 6 offspring, consisting of those litters that produce a total of exactly 6 offspring (i.e., each pair of integers adds up to 6). We say that 6 offspring is a proper subset of Fitness because the elements of the former are completely contained within the latter.

(2,3) (3,3)

(2,4) (4,2)

(2,2) (3,4) (4,3)

(4,4)

(3,2)

6 offspring

Fitness

We can illustrate these two sets with a Venn diagram.12 Figure 1.2 illustrates Venn diagrams for our sets Fitness and 6 offspring. Graphically, you can see that 6 offspring is a proper subset of the larger set Fitness (that is, all of the elements of the former are elements of the latter). We indicate that one set is a subset of another set with the symbol ⊂, and in this case we would write 6 offspring ⊂ Fitness Three of the 9 possible outcomes of Fitness give rise to 6 offspring, and so we would estimate the probability of having 6 offspring to be 1/9 + 1/9 + 1/9 = 3/9 (recalling our assumption that each of the outcomes is equally likely). This result is generalized in the Second Axiom of Probability: Axiom 2: The probability of a complex event equals the sum of the

probabilities of the outcomes that make up that event.

John Venn

John Venn (1834–1923) studied at Gonville and Caius College, Cambridge University, England, from which he graduated in 1857. He was ordained as a priest two years later and returned to Cambridge in 1862 as a lecturer in Moral Science. He also studied and taught logic and probability theory. He is best known for the diagrams that represent sets, their unions and intersections, and these diagrams now bear his name. Venn is also remembered for building a machine for bowling cricket balls.

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The Mathematics of Probability

You can think of a complex event as an or statement in a computer program: if the simple events are A, B, and C, the complex event is (A or B or C), because any one of these outcomes will represent the complex event. Thus P(A or B or C) = P(A) + P(B) + P(C)

(1.2)

For another simple example, consider the probability of drawing a single card from a well-shuffled 52-card deck, and obtaining an ace. We know there are 4 aces in the deck, and the probability of drawing each particular ace is 1/52. Therefore, the probability of the complex event of drawing any 1 of the 4 aces is: P(ace) = 1/52 + 1/52 + 1/52 + 1/52 = 4/52 = 1/13 SHARED EVENTS: MULTIPLYING PROBABILITIES

In the whirligig example, we calculated the probability that a whirligig produces exactly 6 offspring. This complex event could occur by any 1 of 3 different litter pairs {(2,4), (3,3), (4,2)} and we determined the probability of this complex event by adding the simple probabilities. Now we will turn to the calculation of the probability of a shared event—the simultaneous occurrence of two simple events. We assume that the number of offspring produced in the second litter is independent of the number produced in the first litter. Independence is a critical, simplifying assumption for many statistical analyses. When we say that two events are independent of one another, we mean that the outcome of one event is not affected by the outcome of the other. If two events are independent of one another, then the probability that both events occur (a shared event) equals the product of their individual probabilities: P(A ∩ B) = P(A) × P(B) (if A and B are independent)

(1.3)

The symbol ∩ indicates the intersection of two independent events—that is, both events occurring simultaneously. For example, in the litter size example, suppose that an individual can produce 2, 3, or 4 offspring in the first litter, and that the chances of each of these events are 1/3. If the same rules hold for the production of the second litter, then the probability of obtaining the pair of litters (2,4) equals 1/3 × 1/3 = 1/9. Notice that this is the same number we arrived at by treating each of the different litter pairs as independent, equiprobable events. Probability Calculations: Milkweeds and Caterpillars

Here is a simple example that incorporates both complex and shared events. Imagine a set of serpentine rock outcroppings in which we can find populations

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CHAPTER 1 An Introduction to Probability

of milkweed, and in which we also can find herbivorous caterpillars. Two kinds of milkweed populations are present: those that have evolved secondary chemicals that make them resistant (R) to the herbivore, and those that have not. Suppose you census a number of milkweed populations and determine that P(R) = 0.20. In other words, 20% of the milkweed populations are resistant to the herbivore. The remaining 80% of the populations represent the complement of this set. The complement includes all the other elements of the set, which we can write in short hand as not R. Thus P(R) = 0.20

P(not R) = 1 – P(R) = 0.80

Similarly, suppose the probability that the caterpillar (C) occurs in a patch is 0.7: P(C) = 0.7

P(not C) = 1 – P(C) = 0.30

Next we will specify the ecological rules that determine the interaction of milkweeds and caterpillars and then use probability theory to determine the chances of finding either caterpillars, milkweeds, or both in these patches. The rules are simple. First, all milkweeds and all caterpillars can disperse and reach all of the serpentine patches. Milkweed populations can always persist when the caterpillar is absent, but when the caterpillar is present, only resistant milkweed populations can persist. As before, we assume that milkweeds and caterpillars initially colonize patches independently of one another.13 Let’s first consider the different combinations of resistant and non-resistant populations occurring with and without herbivores. These are two simultaneous events, so we will multiply probabilities to generate the 4 possible shared events (Table 1.2). There are a few important things to notice in Table 1.2. First, the sum of the resulting probabilities of the shared events (0.24 + 0.56 + 0.06 + 0.14) = 1.0, and these 4 shared events together form a proper set. Second, we can add some of these probabilities together to define new complex events and also recover some of the underlying simple probabilities. For example, what is the probability that milkweed populations will be resistant? This can be estimated as the probability of finding resistant populations with caterpillars 13

Lots of interesting biology occurs when the assumption of independence is violated. For example, many species of adult butterflies and moths are quite selective and seek out patches with appropriate host plants for laying their eggs. Consequently, the occurrence of caterpillars may not be independent of the host plant. In another example, the presence of herbivores increases the selective pressure for the evolution of host resistance. Moreover, many plant species have so-called facultative chemical defenses that are switched on only when herbivores show up. Consequently, the occurrence of resistant populations may not be independent of the presence of herbivores. Later in this chapter, we will learn some methods for incorporating non-independent probabilities into our calculations of complex events.

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TABLE 1.2 Probability calculations for shared events Outcome Milkweed present?

Caterpillar present?

[1 – P(R)] × [1 – P(C)] = (1.0 – 0.2) × (1.0 – 0.7) = 0.24

Yes

[1 – P(R)] × [P(C)] = (1.0 – 0.2) × (0.7) = 0.56

Yes

[P(R)] × [1 – P(C)] = (0.2) × (1.0 – 0.7) = 0.06

Yes

[P(R)] × [P(C)] = (0.2) × (0.7) = 0.14

Yes

Shared event

Probability calculation

Susceptible population and no caterpillar Susceptible population and caterpillar Resistant population and no caterpillar Resistant population and caterpillar

The joint occurrence of independent events is a shared event, and its probability can be calculated as the product of the probabilities of the individual events. In this hypothetical example, milkweed populations that are susceptible to herbivorous caterpillars occur with probability P(R), and milkweed populations that are resistant to caterpillars occur with probability 1 – P(R). The probability that a milkweed patch is colonized by caterpillars is P(C) and the probability that a milkweed patch is not colonized by caterpillars is 1 – P(C). The simple events are the occurrence of caterpillars (C) and of resistant milkweed populations (R). The first column lists the four complex events, defined by resistant or susceptible milkweed populations and the presence or absence of caterpillars. The second column illustrates the probability calculation of the complex event. The third and fourth columns indicate the ecological outcome. Notice that the milkweed population goes extinct if it is susceptible and is colonized by caterpillars. The probability of this event occurring is 0.56, so we expect this outcome 56% of the time. The most unlikely outcome is a resistant milkweed population that does not contain caterpillars (P = 0.06). Notice also that the four shared events form a proper set, so their probabilities sum to 1.0 (0.24 + 0.56 + 0.06 + 0.14 = 1.0).

(P = 0.14) plus the probability of finding resistant populations without caterpillars (P = 0.06). The sum (0.20) indeed matches the original probability of resistance [P(R) = 0.20]. The independence of the two events ensures that we can recover the original values in this way. However, we have also learned something new from this exercise. The milkweeds will disappear if a susceptible population encounters the caterpillar, and this should happen with probability 0.56. The complement of this event, (1 – 0.56) = 0.44, is the probability that a site will contain a milkweed population. Equivalently, the probability of milkweed persistence can be calculated as P(milkweed present) = 0.24 + 0.06 + 0.14 = 0.44, adding together the probabilities for the different combinations that result in milkweeds. Thus, although the probability of resistance is only 0.20, we expect to find milkweed populations occurring in 44% of the sampled patches because not all susceptible populations are hit by caterpillars. Again, we emphasize that these calculations are correct only if the initial colonization events are independent of one another.

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Complex and Shared Events: Rules for Combining Sets

Many events are not independent of one another, however, and we need methods to take account of that non-independence. Returning to our whirligig example, what if the number of offspring produced in the second litter was somehow related to the number produced in the first litter? This might happen because organisms have a limited amount of energy available for producing offspring, so that energy invested in the first litter of offspring is not available for investment in the second litter. Would that change our estimate of the probability of producing 6 offspring? Before we can answer this question, we need a few more tools in our probabilistic toolkit. These tools tell us how to combine events or sets, and allow us to calculate the probabilities of combinations of events. Suppose in our sample space there are two identifiable events, each of which consists of a group of outcomes. For example, in the sample space Fitness, we could describe one event as a whirligig that produces exactly 2 offspring in its first litter. We will call this event First litter 2, and abbreviate it as F. The second event is a whirligig that produces exactly 4 offspring in its second litter. We will call this event Second litter 4 and abbreviate it as S: Fitness = {(2,2), (2,3), (2,4), (3,2), (3,3), (3,4), (4,2), (4,3), (4,4)} F = {(2,2), (2,3), (2,4)} S = {(2,4), (3,4), (4,4)} We can construct two new sets from F and S. The first is the new set of outcomes that equals all the outcomes that are in either F or S alone. We indicate this new set using the notation F ∪ S, and we call this new set the union of these two sets: F ∪ S = {(2,2), (2,3), (2,4), (3,4), (4,4)} Note that the outcome (2,4) occurs in both F and S, but it is counted only once in F ∪ S. Also notice that the union of F and S is a set that contains more elements than are contained in either F or S, because these sets are “added” together to create the union. The second new set equals the outcomes that are in both F and S. We indicate this set with the notation F ∩ S, and call this new set the intersection of the two sets: F ∩ S = {(2,4)} Notice that the intersection of F and S is a set that contains fewer elements than are contained in either F or S alone, because now we are considering only the elements that are common to both sets. The Venn diagram in Figure 1.3 illustrates these operations of union and intersection.

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Fitness

(2,2) F

(2,3) F∩S

(2,4)

(4,3) (3,3) (3,2)

F∪S

(3,4) (4,4)

(4,2)

Figure 1.3 Venn diagram illustrating unions and intersections of sets. Each ring represents a different set of the numbers of offspring produced in a pair of litters by an imaginary whirligig beetle species, as in Figure 1.2. The largest ring is the set Fitness, which encompasses all of the possible reproductive outputs of the beetle. The small ring F is the set of all pairs of litters in which there are exactly 2 offspring in the first litter. The small ring S is the set of all pairs of litters in which there are exactly 4 offspring in the second litter. The area in which the rings overlap represents the intersection of F and S (F ∩ S) and contains only those elements common to both. The ring that encompasses F and S represents the union of F and S (F ∪ S) and contains all elements found in either set. Notice that the union of F and S does not double count their common element, (2,4). In other words, the union of two sets is the sum of the elements in both sets, minus their common elements. Thus, F ∪ S = (F + S) – (F ∩ S).

We can construct a third useful set by considering the set F c, called the complement of F, which is the set of objects in the remaining sample space (in this example, Fitness) that are not in the set F: F c = {(3,2), (3,3), (3,4), (4,2), (4,3), (4,4)} From Axioms 1 and 2, you should see that P(F) + P(F c) = 1.0 In other words, because F and F c collectively include all possible outcomes (by definition, they comprise the entire set), the sum of the probabilities associated with them must add to 1.0. Finally, we need to introduce the empty set. The empty set contains no elements and is written as {∅}. Why is this set important? Consider the set consisting of the intersection of F and F c. Because they have no elements in common, if we did not have an empty set, then F ∩ F c would be undefined. Having

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an empty set allows sets to be closed14 under the three allowable operations: union, intersection, and complementation. CALCULATING PROBABILITIES OF COMBINED EVENTS Axiom 2 stated that the probability of a complex event equals the sum of the probabilities of the outcomes that make up that event. It should be a simple operation, therefore, to calculate the probability of, say, F ∪ S as P(F) + P(S). Because both F and S have 3 outcomes, each with probability 1/9, this simple sum gives a value of 6/9. However, there are only 5 elements in F ∪ S, so the probability should be only 5/9. Why didn’t the axiom work? Figure 1.3 showed that these two sets have the outcome (2,4) in common. It is no coincidence that this outcome is equal to the intersection of F and S. Our simple calculation of the union of these two sets resulted in our counting this common element twice. In general, we need to avoid double counting when we calculate the probability of a union. 15

Closure is a mathematical property, not a psychological state. A collection of objects G (technically, a group) is closed under an operation ⊕ (such as union, intersection, or complementation) if, for all elements A and B of G, A ⊕ B is also an element of G.

Determining whether events have been double-counted or not can be tricky. For example, in basic population genetics, the familiar Hardy-Weinberg equation gives the frequencies of different genotypes present in a randomly mating population. For a single-gene, two-allele system, the Hardy-Weinberg equation predicts the frequency of each genotype (RR, Rr, and rr) in a population in which p is the initial allele frequency of R and q is the initial allele frequency of r. Because the genotypes R and r form a closed set, p + q = 1.0. Each parent contributes a single allele in its gametes, so the formation of offspring represents a shared event, with two gametes combining at fertilization and contributing their alleles to determine the genotype of the offspring. The Hardy-Weinberg equation predicts the probability (or frequency) of the different genotype combinations in the offspring of a randomly mating population as P(RR) = p2 P(Rr) = 2pq P(rr) = q2

The production of an RR genotype requires that both gametes contain the R allele. Because the frequency of the R allele in the population is p, the probability of this complex event = p × p = p2. But why does the heterozygote genotype (Rr) occur with frequency 2pq and not just pq? Isn’t this a case of double counting? The answer is no, because there are two ways to create a heterozygous individual: we can combine a male R gamete with a female r gamete, or we can combine a male r gamete with a female R gamete. These are two distinct events, even though the resulting zygote has an identical genotype (Rr) in both cases. Therefore, the probability of heterozygote formation is a complex event, whose elements are two shared events: P(Rr) = P(male R and female r) or P(male r and female R) P(Rr) = pq + qp = 2pq

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We therefore calculate the probability of a union of any two sets or events A and B as P(A ∪ B) = P(A) + P(B) – P(A ∩ B)

(1.4)

The probability of an intersection is the probability of both events happening. In the example shown in Figure 1.3, F ∩ S = {(2,4)}, which has probability 1/9, the product of the probabilities of F and S ((1/3) × (1/3) = 1/9). So, subtracting 1/9 from 6/9 gives us the desired probability of 5/9 for P(F ∪ S). If there is no overlap between A and B, then they are mutually exclusive events, meaning they have no outcomes in common. The intersection of two mutually exclusive events (A ∩ B) is therefore the empty set {∅}. Because the empty set has no outcomes, it has probability = 0, and we can simply add the probabilities of two mutually exclusive events to obtain the probability of their union: P(A ∪ B) = P(A) + P(B)

(if A ∩ B = {∅})

(1.5)

We are now ready to return to our question of estimating the probability that an orange-spotted whirligig producing 6 offspring, if the number of offspring produced in the second litter depends on the number of offspring produced in the first litter. Recall our complex event 6 offspring, which consisted of the three outcomes (2,4), (3,3), and (4,2); its probability is P(6 offspring) = 3/9 (or 1/3). We also found another set F, the set for which the number of offspring in the first litter was equal to 2, and we found that P(F) = 3/9 (or 1/3). If you observed that the first litter was 2 offspring, what is the probability that the whirligig will produce 4 offspring the next time (for a total of 6)? Intuitively, it seems that this probability should be 1/3 as well, because there are 3 outcomes in F, and only one of them (2,4) results in a total of 6 offspring. This is the correct answer. But why is this probability not equal to 1/9, which is the probability of getting (2,4) in Fitness? The short answer is that the additional information of knowing the size of the first litter influenced the probability of the total reproductive output. Conditional probabilities resolve this puzzle. Conditional Probabilities

If we are calculating the probability of a complex event, and we have information about an outcome in that event, we should modify our estimates of the probabilities of other outcomes accordingly. We refer to these updated estimates as conditional probabilities. We write this as P(A | B) or the probability of event or outcome A given event or outcome B. The vertical slash (|) indicates that the probability of A is calculated assuming that the

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event or outcome B has already occurred. This conditional probability is defined as P (A | B) =

P (A ∩ B) P (B)

(1.6)

This definition should make intuitive sense. If outcome B has already occurred, then any outcomes in the original sample space not in B (i.e., B c) cannot occur, so we restrict the outcomes of A that we could observe to those that also occur in B. So, P(A | B) should somehow be related to P(A ∩ B), and we have defined it to be proportional to that intersection. The denominator P(B) is the restricted sample space of events for which B has already occurred. In our whirligig example, P(F ∩ S) = 1/9, and P(F) = 1/3. Dividing the former by the latter gives the value for P(F | S) = (1/9)/(1/3) = 1/3, as suggested by our intuition. Rearranging the formula in Equation 1.6 for calculating conditional probabilities gives us a general formula for calculating the probability of an intersection: P(A ∩ B) = P(A | B) × P(B) = P(B | A) × P(A)

(1.7)

You should recall that earlier in the chapter we defined the probability of the intersection of two independent events to be equal to P(A) × P(B). This is a special case of the formula for calculating the probability of intersection using the conditional probability P(A | B). Simply note that if two events A and B are independent, then P(A | B) = P(A), so that P(A | B) × P(B) = P(A) × P(B). By the same reasoning, P(A ∩ B) = P(B | A) × P(A).

Bayes’ Theorem Until now, we have discussed probability using what is known as the frequentist paradigm, in which probabilities are estimated as the relative frequencies of outcomes based on an infinitely large set of trials. Each time scientists want to estimate the probability of a phenomenon, they start by assuming no prior knowledge of the probability of an event, and re-estimate the probability based on a large number of trials. In contrast, the Bayesian paradigm, based on a formula for conditional probability developed by Thomas Bayes,16 builds on the 16

Thomas Bayes

Thomas Bayes (1702–1761) was a Nonconformist minister in the Presbyterian Chapel in Tunbridge Wells, south of London, England. He is best known for his “Essay towards solving a problem in the doctrine of chances,” which was published two years after his death in Philosophical Transactions 53: 370–418 (1763). Bayes was elected a Fellow of the Royal Society of London in 1742. Although he was cited for his contributions to mathematics, he never published a mathematical paper in his lifetime.

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Bayes’ Theorem

idea that investigators may already have a belief of the probability of an event, before the trials are conducted. These prior probabilities may be based on previous experience (which itself could be a frequentist estimate from earlier studies), intuition, or model predictions. These prior probabilities are then modified by the data from the current trials to yield posterior probabilities (discussed in more detail in Chapter 5). However, even in the Bayesian paradigm, quantitative estimates of prior probabilities will ultimately have to come from trials and experiments. Bayes’ formula for calculating conditional probabilities, now known as Bayes’ Theorem, is P( A | B) =

P(B | A)P( A) P(B | A)P( A) + P(B | Ac )P ( Ac )

(1.8)

This formula is obtained by simple substitution. From Equation 1.6, we can write the conditional probability P(A | B) on the right side of the equation as P( A | B) =

P( A ∩ B) P(B)

From Equation 1.7, the numerator of this second equation can be rewritten as P(B | A) × P(A), yielding the numerator of Bayes’ Theorem. By Axiom 1, the denominator, P(B), can be rewritten as P(B ∩ A) + P(B ∩ Ac), as these two terms sum to P(B). Again using our formula for the probability of an intersection (Equation 1.7), this sum can be rewritten as P(B | A) × P(A) + P(B | Ac) × P (Ac), which is the denominator of Bayes’ Theorem. Although Bayes’ Theorem is simply an expansion of the definition of conditional probability, it contains a very powerful idea. That idea is that the probability of an event or outcome A conditional on another event B can be determined if you know the probability of the event B conditional on the event A and you know the complement of A, Ac (which is why Bayes’ Theorem is often called a theorem of inverse probability). We will return to a detailed exploration of Bayes’ Theorem and its use in statistical inference in Chapter 5. For now, we conclude by highlighting the very important distinction between P(A | B) and P(B | A). Although these two conditional probabilities look similar, they are measuring completely different things. As an example, let’s return to the susceptible and resistant milkweeds and the caterpillars. First consider the conditional probability P(C | R) This expression indicates the probability that caterpillars (C) are found given a resistant population of milkweeds (R). To estimate P(C | R), we would need to

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examine a random sample of resistant milkweed populations and to determine the frequency with which these populations were hosting caterpillars. Now consider this conditional probability: P(R | C) In contrast to the previous expression, P(R | C) is the probability that a milkweed population is resistant (R), given that it is being eaten by caterpillars (C). To estimate P(R | C), we would need to examine a random sample of caterpillars and determine the frequency with which they were actually feeding on resistant milkweed populations. These are two quite different quantities and their probabilities are determined by different factors. The first conditional probability P(C | R) (the probability of the occurrence of caterpillars given resistant milkweeds) will depend, among other things, on the extent to which resistance to herbivory is directly or indirectly responsible for the occurrence of caterpillars. In contrast, the second conditional probability P(R | C) (the probability of resistant milkweed populations given the presence of caterpillars) will depend, in part, on the incidence of caterpillars on resistant plants versus other situations that could also lead to the presence of caterpillars (e.g., caterpillars feeding on other plant species or caterpillars feeding on susceptible plants that were not yet dead). Finally, note that the conditional probabilities are also distinct from the simple probabilities: P(C) is just the probability that a randomly chosen individual is a caterpillar, and P(R) is the probability that a randomly chosen milkweed population is resistant to caterpillars. In Chapter 5, we will see how to use Bayes’ Theorem to calculate conditional probabilities when we cannot directly measure P(A | B).

Summary The probability of an outcome is simply the number of times that the outcome occurs divided by the total number of trials. If simple probabilities are known or estimated, probabilities can be determined for complex events (Event A or Event B) through summation, and probabilities can be determined for shared events (Event A and Event B) through multiplication. The definition of probability, together with axioms for the additivity of probabilities and three operations on sets (union, intersection, complementation), form the fundamentals of the probability calculus.

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CHAPTER 2

Random Variables and Probability Distributions

In Chapter 1, we explored the notion of probability and the idea that the outcome of a single trial is an uncertain event. However, when we accumulate data on many trials, we may begin to see regular patterns in the frequency distribution of events (e.g., Figure 1.1). In this chapter, we will explore some useful mathematical functions that can generate these frequency distributions. Certain probability distributions are assumed by many of our common statistical tests. For example, parametric analysis of variance (ANOVA; see Chapter 10) assumes that random samples of measured values fit a normal, or bellshaped, distribution, and that the variance of that distribution is similar among different groups. If the data meet those assumptions, ANOVA can be used to test for differences among group means. Probability distributions also can be used to build models and make predictions. Finally, probability distributions can be fit to real datasets without specifying a particular mechanistic model. We use probablility distributions because they work—they fit lots of data in the real world. Our first foray into probability theory involved estimating the probabilities of individual outcomes, such as prey capture by pitcher plants or reproduction of whirligig beetles. We found numerical values for each of these probabilities by using simple rules, or functions. More formally, the mathematical rule (or function) that assigns a given numerical value to each possible outcome of an experiment in the sample space of interest is called a random variable. We use the term “random variable” in this mathematical sense, not in the colloquial sense of a random event. Random variables come in two forms: discrete random variables and continuous random variables. Discrete random variables are those that take on finite or countable values (such as integers; see Footnote 4 in Chapter 1). Common examples include presence or absence of a given species (which takes the

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value of 1 or 0), or number of offspring, leaves, or legs (integer values). Continuous random variables, on the other hand, are those that can take on any value within a smooth interval. Examples include the biomass of a starling, the leaf area consumed by an herbivore, or the dissolved oxygen content of a water sample. In one sense, all values of a continuous variable that we measure in the real world are discrete: our instrumentation only allows us to measure things with a finite level of precision.1 But there is no theoretical limit to the precision we could obtain in the measurement of a continuous variable. In Chapter 7, we will return to the distinction between discrete and continuous variables as a key consideration in the design of field experiments and sampling designs.

Discrete Random Variables Bernoulli Random Variables

The simplest experiment has only two outcomes, such as organisms being present or absent, coins landing heads or tails, or whirligigs reproducing or not. The random variable describing the outcome of such an experiment is a Bernoulli random variable, and an experiment of independent trials in which there are only two possible outcomes for each trial is a Bernoulli trial.2 We use the notation 1

It is important to understand the distinction between precision and accuracy in measurements. Accuracy refers to how close the measurement is to the true value. Accurate measurements are unbiased, meaning they are neither consistently above nor below the true value. Precision refers to the agreement among a series of measurements and the degree to which these measurements can be discriminated. For example, a measurement can be precise to 3 decimal places, meaning we can discriminate among different measurements out to 3 decimal places. Accuracy is more important than precision. It would be much better to use an accurate balance that was precise to only 1 decimal place than an inaccurate balance that was precise to 5 decimal places. The extra decimal places don’t bring you any closer to the true value if your instrument is flawed or biased. 2

In our continuing series of profiles of dead white males in wigs, Jacob (aka Jaques or James) Bernoulli (1654–1705) was one of a family of famous physicists and mathematicians. He and his brother Johann are considered to be second only to Newton in their development of calculus, but they argued constantly about the relative merits of each other’s work. Jacob’s major mathematical work, Ars conjectandi (published in 1713, 8 years after his death), was the first major text on probability. It included the first exposition of many of Jacob Bernoulli the topics discussed in Chapters 2 and 3, such as general theories of permutations and combinations, the first proofs of the binomial theorem, and the Law of Large Numbers. Jacob Bernoulli also made substantial contributions to astronomy and mechanics (physics).

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Discrete Random Variables

X ∼ Bernoulli(p)

(2.1)

to indicate that the random variable X is a Bernoulli random variable. The symbol ∼ is read “is distributed as.” X takes on the values of the number of “successes” in the trial (e.g., present, captured, reproducing), and p is the probability of a successful outcome. The most common example of a Bernoulli trial is a flip of a single fair coin (perhaps not a Belgian Euro; see Footnote 7 in Chapter 1, and Chapter 11), where the probability of heads = the probability of tails = 0.5. However, even a variable with a large number of outcomes can be redefined as a Bernoulli trial if we can collapse the range of responses to two outcomes. For example, a set of 10 reproductive events of whirligig beetles can be analyzed as a single Bernoulli trial, where success is defined as having exactly 6 offspring and failure is defined as having more than or fewer than 6 offspring. An Example of a Bernoulli Trial

A one-time census of all the towns in Massachusetts for the rare plant species Rhexia mariana (meadow beauty) provides an example of a Bernoulli trial. The occurrence of Rhexia is a Bernoulli random variable X, which takes on two outcomes: X = 1 (Rhexia present) or X = 0 (Rhexia absent). There are 349 towns in Massachusetts, so our single Bernoulli trial is the one search of all of these towns for occurrences of Rhexia. Because Rhexia is a rare plant, let us imagine that the probability that Rhexia is present (i.e., X = 1) is low: P(X = 1) = p = 0.02. Thus, if we census a single town, there is only a 2% chance (p = 0.02) of finding Rhexia. But what is the expected probability that Rhexia occurs in any 10 towns, and not in any of the remaining 339 towns? Because the probability of Rhexia being present in any given town p = 0.02, we know from the First Axiom of Probability (see Chapter 1) that the probability of Rhexia being absent from any single town = (1 − p) = 0.98. By definition, each event (occurrence in a single town) in our Bernoulli trial must be independent. For this example, we assume the presence or absence of Rhexia in a given town to be independent of its presence or absence in any other town. Because the probability that Rhexia occurs in one town is 0.02, and the probability that it occurs in another town is also 0.02, the probability that it occurs in both of these specific towns is 0.02 × 0.02 = 0.0004. By extension, the probability that Rhexia occurs in 10 specified towns is 0.0210 = 1.024 × 10–17 (or 0.00000000000000001024, which is a very small number). However, this calculation does not give us the exact answer we are looking for. More precisely, we want the probability that Rhexia occurs in exactly 10 towns and does not occur in the remaining 339 towns. Imagine that the first town surveyed has no Rhexia, which should occur with probability (1 − p) = 0.98. The second town has Rhexia, which should occur with probability p = 0.02. Because the

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presence or absence of Rhexia in each town is independent, the probability of getting one town with Rhexia and one without by picking any two towns at random should be the product of p and (1 − p) = 0.02 × 0.98 = 0.0196 (see Chapter 1). By extension, then, the probability of Rhexia occurring in a given set of 10 towns in Massachusetts should be the probability of occurrence in 10 towns (0.0210) multiplied by the probability that it does not occur in the remaining 339 towns (0.98339) = 1.11 × 10 –20. Once again this is an astronomically small number. But we are not finished yet! Our calculation so far gives us the probability that Rhexia occurs in exactly 10 particular towns (and in no others). But we are actually interested in the probability that Rhexia occurs in any 10 towns in Massachusetts, not just a specific list of 10 particular towns. From the Second Axiom of Probability (see Chapter 1), we learned that probabilities of complex events that can occur by different pathways can be calculated by adding the probabilities for each of the pathways. So, how many different sets of 10 town each are possible from a list of 349? Lots! In fact, 6.5 × 1018 different combinations of 10 are possible in a set of 349 (we explain this calculation in the next section). Therefore, the probability that Rhexia occurs in any 10 towns equals the probability that it occurs in one set of 10 towns (0.0210) times the probability that it does not occur in the remaining set of 339 towns (0.98339) times the number of unique combinations of 10 towns that can be produced from a list of 349 (6.5 × 1018). This final product = 0.07; there is a 7% chance of finding exactly 10 towns with Rhexia. This is also a small number, but not nearly as small as the first two probability values we calculated. Many Bernoulli Trials = A Binomial Random Variable

Because a central feature of experimental science is replication, we would rarely conduct a single Bernoulli trial. Rather, we would conduct replicate, independent Bernoulli trials in a single experiment. We define a binomial random variable X to be the number of successful results in n independent Bernoulli trials. Our shorthand notation for a binomial random variable is X ∼ Bin(n,p)

(2.2)

to indicate the probability of obtaining X successful outcomes in n independent Bernoulli trials, where the probability of a successful outcome of any given event is p. Note that if n = 1, then the binomial random variable X is equivalent to a Bernoulli random variable. Binomial random variables are one of the most common types of random variables encountered in ecological and environmental studies. The probability of obtaining X successes for a binomial random variable is

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Discrete Random Variables

P( X ) =

n! p X (1 − p)n− X X !(n − X )!

(2.3)

where n is the number of trials, X is the number of successful outcomes (X ≤ n), and n! means n factorial,3 which is calculated as n × (n−1) × (n−2) × … × (3) × (2) × (1) Equation 2.3 has three components, and two of them should look familiar from our analysis of the Rhexia problem. The component pX is the probability of obtaining X independent successes, each with probability p. The component (1 – p)(n–X) is the probability of obtaining (n – X) failures, each with probability (1 – p). Note that the sum of the successes (X) and the failures (n – X) is just n, the total number of Bernoulli trials. As we saw in the Rhexia example, the probability of obtaining X successes with probability p and (n – X) failures with probability (1 – p) is the product of these two independent events p X(1 – p)(n–X). But then why do we need the term n! X !(n − X )! and where does it come from? The equivalent notation for this term is ⎛ n⎞ ⎜ ⎟ ⎝X⎠ (read as “n choose X”), which is known as the binomial coefficient. The binomial coefficient is needed because there is more than one way to obtain most combinations of successes and failures (the combinations of 10 towns we described in the Rhexia example above). For example, the outcome one success in a set of two Bernoulli trials can actually occur in two ways: (1,0) or (0,1). So the probability of getting one success in a set of two Bernoulli trials equals the probability of an outcome of one success [= p(1 – p)] times the number of possible outcomes of one success (= 2). We could write out all of the different outcomes with X successes and count them, but as n gets large, so does X; there are 2n possible outcomes for n trials. It is more straightforward to compute directly the number of outcomes of X (successes), and this is what the binomial coefficient does. Returning to the Rhexia example, we have 10 occurrences of this plant (X = 10) and 349 towns (n = 349), but we don’t know which particular towns they are in. Beginning our

The factorial operation can be applied only to non-negative integers. By definition, 0! = 1.

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search, there initially are 349 towns in which Rhexia could occur. Once Rhexia is found in a given town, there are only 348 other towns in which it could occur. So there are 349 combinations of towns in which you could have only one occurrence, and 348 combinations in which you could have the second occurrence. By extension, for X occurrences, the total number of ways that 10 Rhexia occurrences could be distributed among the 349 towns would be 349 × 348 × 347 × 346 × 345 × 344 × 343 × 342 × 341 × 340 = 2.35 × 1025 In general, the total number of ways to obtain X successes in n trials is n × (n – 1) × (n – 2) × … × (n – X + 1) which looks a lot like the formula for n!. The terms in the above equation that are missing from n! are all the remaining ones below (n – X + 1), or (n – X) × (n – X – 1) × (n – X – 2) × … × 1 which simply equals (n – X)!. Thus, if we divide n! by (n – X)!, we’re left with the total number of ways of obtaining X successes in n trials. But this is not quite the same as the binomial coefficient described above, which further divides our result by X!: n! (n − X )! X ! The reason for the further division is that we don’t want to double-count identical patterns of successes that simply occurred in a different order. Returning to Rhexia, if we found populations first in the town of Barnstable and second in Chatham, we would not want to count that as a different outcome from finding it first in Chatham and second in Barnstable. By the same reasoning as we used above, there are exactly X! ways of reordering a given outcome. Thus, we want to “discount” (divide by) our result by X!. The utility of such different permutations will become more apparent in Chapter 5, when we discuss Monte Carlo methods for hypothesis testing.4

One more example to persuade yourself that the binomial coefficient works. Consider the following set of five marine fishes: {(wrasse), (blenny), (goby), (eel), (damselfish)} How many unique pairs of fishes can be formed? If we list all of them, we find 10 pairs: (wrasse), (blenny) (wrasse), (goby) (wrasse), (eel) (wrasse), (damselfish) (blenny), (goby)

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The Binomial Distribution

Now that we have a simple function for calculating the probability of a binomial random variable, what do we do with it? In Chapter 1, we introduced the histogram, a type of graph used to summarize concisely the number of trials that resulted in a particular outcome. Similarly, we can plot a histogram of the number of binomial random variables from a series of Bernoulli trials that resulted in each possible outcome. Such a histogram is called a binomial distribution, and it is generated from a binomial random variable (Equation 2.3). For example, consider the Bernoulli random variable for which the probability of success = the probability of failure = 0.5 (such as flipping fair coins). Our experiment will consist of flipping a fair coin 25 times (n = 25), and our possible outcomes are given by the sample space number of heads = {0, 1, …, 24, 25}. Each outcome Xi is a binomial random variable, and the probability of each outcome is given by the binomial formula, which we will refer to as a probability distribution function, because it is the function (or rule) that provides the numerical value (the probability) of each outcome in the sample space. Using our formula for a binomial random variable, we can tabulate the probabilities of every possible outcome from hunting for Rhexia 25 times (Table 2.1). We can draw this table as a histogram (Figure 2.1), and the histogram can be interpreted in two different ways. First, the values on the y-axis can be read as the probability of obtaining a given random variable X (e.g., the number of occurrences of Rhexia) in 25 trials where the probability of an occurrence is 0.5. In this interpretation, we refer to Figure 2.1 as a probability distribution. Accordingly, if we add up the values in Table 2.1, they sum to exactly 1.0 (with rounding error), because they define the entire sample space (from the First Axiom of Probability). Second, we can interpret the values on the y-axis as the expected relative frequency of each random variable in a large number of experiments, each of which had 25 replicates. This definition of relative frequency matches the formal definition of probability given in Chapter 1. It is also the basis of the term frequentist to describe statistics based on the expected frequency of an event based on an infinitely large number of trials. (blenny), (eel) (blenny), (damselfish) (goby), (eel) (goby), (damselfish) (eel), (damselfish) Using the binomial coefficient, we would set n = 5 and X = 2 and arrive at the same number: ⎛ 5⎞ 5! 120 ⎜ ⎟ = 3! 2! = 6 × 2 = 10 ⎝ 2⎠

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The first column gives the full range of possible numbers of successes out of 25 trials (i.e., from 0 to 25). The second column gives the probability of obtaining exactly that number of successes, calculated from Equation 2.3. Allowing for rounding error, these probabilities sum to 1.0, the total area under the probability curve. Notice that a binomial with p = 0.5 gives a perfectly symmetric distribution of successes, centered around 12.5, the expectation of the distribution (Figure 2.1).

TABLE 2.1 Binomial probabilities for p = 0.5, 25 trials Number of successes (X)

0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25

Probability of X in 25 trials (P(X))

0.00000003 0.00000075 0.00000894 0.00006855 0.00037700 0.00158340 0.00527799 0.01432598 0.03223345 0.06088540 0.09741664 0.13284087 0.15498102 0.15498102 0.13284087 0.09741664 0.06088540 0.03223345 0.01432598 0.00527799 0.00158340 0.00037700 0.00006855 0.00000894 0.00000075 0.00000003 Σ = 1.00000000

Figure 2.1 is symmetric—the left- and right-hand sides of the distribution are mirror images of one another.5 However, that result is a special property of a binomial distribution for which p = (1 – p) = 0.50. Alternatives are possible, 5

If you ask most people what the chances are of obtaining 12 or 13 heads out of 25 tosses of a fair coin, they will say about 50%. However, the actual probability of obtaining 12 heads is only 0.155 (Table 2.1), about 15.5%. Why is this number so small? The answer is that the binomial equation gives the exact probability: the value of 0.155 is the probability of obtaining exactly 12 heads—no more or less. However, scientists are

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Discrete Random Variables

0.16 X ~ Bin(25, 0.5)

P(X)

0.12

0.08

0.04

0.00 0

Figure 2.1 Probability distribution for a binomial random variable. A binomial random variable has only two possible outcomes (e.g., Yes, No) in a single trial. The variable X represents the number of positive outcomes of 25 trials. The variable P(X) represents the probability of obtaining that number of positive outcomes, calculated from Equation 2.3. Because the probability of a successful outcome was set at p = 0.5, this binomial distribution is symmetric, and the midpoint of the distribution is 12.5, or half of the 25 trials. A binomial distribution is specified by two parameters: n, the number trials; and p, the probability of a positive outcome.

such as with biased coins, which come up heads more frequently than 50% of the time. For example, setting p = 0.80 and 25 trials, a different shape of the binomial distribution is obtained, as shown in Figure 2.2. This distribution is asymmetric and shifted toward the right; samples with many successes are more probable than they are in Figure 2.1. Thus, the exact shape of the binomial distribution depends both on the total number of trials n, and the probability of sucoften more interested in knowing the extreme or tail probability. The tail probability is the chance of obtaining 12 or fewer heads in 25 trials. The tail probability is calculated by adding the probabilities for each of the outcomes from 0 to 12 heads. This sum is indeed 0.50, and it corresponds to the area under the left half of the binomial distribution in Figure 2.1.

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0.20

X ~ Bin(25, 0.8)

0.15

P(X)

0.10

0.05

0.00 0

Figure 2.2 Probability distribution for a binomial random variable X ~ Bin(25, 0.8)—that is, there are 25 trials, and the probability of a successful outcome in each trial is p = 0.8. This figure follows the same layout and notation as Figure 2.1. However, here the probability of a positive outcome is p = 0.8 (instead of p = 0.5), and the distribution is no longer symmetric. Notice the expectation for this binomial distribution is 25 × 0.8 = 20, which corresponds to the mode (highest peak) of the histogram.

cess, p. You should try generating your own binomial distributions6 using different values for n and p. Poisson Random Variables

The binomial distribution is appropriate for cases in which there is a fixed number of trials (n) and the probability of success is not too small. However, the formula quickly becomes cumbersome when n becomes large and p becomes small, such as the occurrences of rare plants or animals. Moreover, the binomial is only 6

In Chapter 1 (see Footnote 7), we discussed a coin-spinning experiment for the new Euro coin, which yielded an estimate of p = 0.56 for n = 250 trials. Use spreadsheet or statistical software to generate the binomial distribution with these parameters, and compare it to the binomial distribution for a fair coin (p = 0.50, n = 250). You will see that these probability distributions do, indeed differ, and that the p = 0.56 is shifted very slightly to the right. Now try the same exercise with n = 25, as in Figure 2.1. With this relatively small sample size, it will be virtually impossible to distinguish the two distributions. In general, the closer two distributions are in their expectation (see Chapter 3), the larger a sample we will need to distinguish between them.

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useful when we can directly count the trials themselves. However, in many cases, we do not count individual trials, but count events that occur within a sample. For example, suppose the seeds from an orchid (a flowering plant) are randomly scattered in a region, and we count the number of seeds in sampling quadrats of fixed size. In this case, the occurrence of each seed represents a successful outcome of an unobserved dispersal trial, but we really cannot say how many trials have taken place to generate this distribution. Samples in time can also be treated this way, such as the counts of the number of birds visiting a feeder over a 30-minute period. For such distributions, we use the Poisson distribution rather than the binomial distribution. A Poisson random variable7 X is the number of occurrences of an event recorded in a sample of fixed area or during a fixed interval of time. Poisson random variables are used when such occurrences are rare—that is, when the most common number of counts in any sample is 0. The Poisson random variable itself is the number of events in each sample. As always, we assume that the occurrences of each event are independent of one another. Poisson random variables are described by a single parameter λ, sometimes referred to as a rate parameter because Poisson random variables can describe the frequency of rare events in time. The parameter λ is the average value of the number of occurrences of the event in each sample (or over each time interval). Estimates of λ can be obtained either from data collection or from prior knowledge. Our shorthand notation for a Poisson random variable is X ∼ Poisson(λ)

(2.4)

7 Poisson random variables are named for Siméon-Denis Poisson (1781–1840), who claimed that “life was only good for two things: to do mathematics and to teach it” (fide Boyer 1968). He applied mathematics to physics (in his two-volume work Traité de mécanique, published in 1811 and 1833), and provided an early derivation of the Law of Large Numbers (see Chapter 3). His treatise on probability, Recherches sur la probabilité des jugements, was published in 1837. Siméon-Denis Poisson The most famous literary reference to the Poisson distribution occurs in Thomas Pynchon’s novel Gravity’s Rainbow (1972). One of the lead characters of the novel, Roger Mexico, works for The White Visitation in PISCES (Psychological Intelligence Scheme for Expediting Surrender) during World War II. He plots on a map of London the occurrence of points hit by Nazi bombs, and fits the data to a Poisson distribution. Gravity’s Rainbow, which won the National Book Award in 1974, is filled with other references to statistics, mathematics, and science. See Simberloff (1978) for a discussion of entropy and biophysics in Pynchon’s novels.

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and we calculate the probability of any observation x as P (x ) =

λx – λ e x!

(2.5)

where e is a constant, the base of the natural logarithm (e ∼ 2.71828). For example, suppose the average number of orchid seedlings found in a 1-m2 quadrat is 0.75. What are the chances that a single quadrat will contain 4 seedlings? In this case, λ = 0.75 and x = 4. Using Equation 2.5, P(4 seedlings) =

0.754 –0.75 e = 0.0062 4!

A much more likely event is that a quadrat will contain no seedlings: P(0 seedlings) =

0.750 –0.75 e = 0.4724 0!

There is a close relationship between the Poisson and the binomial distributions. For a binomial random variable X ∼ Bin(n, p), where the number of successes X is very small relative to the sample size (or number of trials) n, we can approximate X by using the Poisson distribution, and by estimating P(X) as Poisson(λ). However, the binomial distribution depends on both the probability of success p and the number of trials n, whereas the Poisson distribution depends only on the average number of events per sample, λ. An entire family of Poisson distributions is possible, depending on the value of λ (Figure 2.3). When λ is small, the distribution has a strong “reverse-J” shape, with the most likely events being 0 or 1 occurrences per sample, but with a long probability tail extending to the right, corresponding to very rare samples that contain many events. As λ increases, the center of the Poisson distribution shifts to the right and becomes more symmetric, resembling a normal or binomial distribution. The binomial and the Poisson are both discrete distributions that take on only integer values, with a minimum value of 0. However, the binomial is always bounded between 0 and n (the number of trials). In contrast, the right-hand tail of the Poisson is not bounded and extends to infinity, although the probabilities quickly become vanishingly small for large numbers of events in a single sample. An Example of a Poisson Random Variable: Distribution of a Rare Plant

To close this discussion, we describe the application of the Poisson distribution to the distribution of the rare plant Rhexia mariana in Massachusetts. The data consist of the number of populations of Rhexia recorded in each of 349 Massachusetts towns between 1913 and 2001 (Craine 2002). Although each population can be thought of as an independent Bernoulli trial, we know only the num-

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λ = 0.1

P(X)

0.9

0.6

0.4

0.3

0.2

0.0

0.4

λ = 1.0

0.3

λ = 0.5

0.3

λ = 2.0

0.2

0.2 0.1 0.1 0.0

0.0 0

6 X

12 0.3

λ = 4.0

0.2

0.1

0.0 0

6 X

Figure 2.3 Family of Poisson curves. Each histogram represents a probability distribution with a different rate parameter λ. The value of X indicates the number of positive outcomes, and P(X) is the probability of that outcome, calculated from Equation 2.5. In contrast to the binomial distributions shown in Figures 2.1 and 2.2, the Poisson distribution is specified by only a single parameter, λ, which is the rate of occurrence of independent events (or the number that occur during some specified time interval). As the value of λ increases, the Poisson distribution becomes more symmetric. When λ is very large, the Poisson distribution is almost indistinguishable from a normal distribution.

ber of populations, not the number of individual plants; the data from each town is our sampling unit. For these 349 sampling units (towns), most samples (344) had 0 populations in any year, but one town has had as many as 5 populations. The average number of populations is 12/349, or 0.03438 populations/town, which we will use as the rate parameter λ in a Poisson distribution. Next, we use

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Equation 2.5 to calculate the probability of a given number of populations in each town. Multiplying those probabilities by the total sample size (n = 349) gives the expected frequencies in each population class (0 through 5), which can be compared to the observed numbers (Table 2.2). The observed frequencies of populations match very closely the predictions from the Poisson model; in fact, they are not significantly different from each other (we used a chi-square test to compare these distributions; see Chapter 11). We can imagine that the town that has five populations of this rare plant is a key target for conservation and management activities—is the soil especially good there, or are there unique plant habitats in that town? Regardless of the ultimate cause of Rhexia occurrence and persistence in these towns, the close match of the data with the Poisson distribution suggests that the populations are random and independent; the five populations in a single town may represent nothing more or less than good luck. Therefore, establishing management strategies by searching for towns with the same set of physical conditions may not result in the successful conservation of Rhexia in those towns.

TABLE 2.2 Expected and observed frequencies of numbers of populations of a rare plant (Rhexia mariana, meadow beauty) in Massachusetts towns Number of Rhexia populations

Poisson probability

Poisson expected frequency

Observed frequency

0 1 2 3 4 5 Total

0.96 0.033 5.7 × 10–4 6.5 × 10–6 5.6 × 10–8 3.8 × 10–10 1.0000

337.2 11.6 0.19 2.3 × 10–3 1.9 × 10–5 1.4 × 10–7 349

344 3 0 0 1 1 349

The first column gives the number of populations found in a particular number of towns. Although there is no theoretical maximum for this number, the observed number was never greater than 5, so only the values for 0 through 5 are shown. The second column gives the Poisson probability for obtaining the observed number, assuming a Poisson distribution with a mean rate parameter λ = 0.03438 populations/town. This value corresponds to the observed frequency in the data (12 observed populations divided by 349 censused towns = 0.03438). The third column gives the expected Poisson frequency, which is simply the Poisson probability multiplied by the total sample size of 349. The final column gives the observed frequency, that is, the number of towns that contained a certain number of Rhexia populations. Notice the close match between the observed and expected Poisson frequencies, suggesting that the occurrence of Rhexia populations in a town is a random, independent event. (Data from Craine 2002.)

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The Expected Value of a Discrete Random Variable

Random variables take on many different values, but the entire distribution can be summarized by determining the typical, or average, value. You are familiar with the arithmetic mean as the measure of central tendency for a set of numbers, and in Chapter 3 we will discuss it and other useful summary statistics that can be calculated from data. However, when we are dealing with probability distributions, a simple average is misleading. For example, consider a binomial random variable X that can take on the values 0 and 50 with probabilities 0.1 and 0.9, respectively. The arithmetic average of these two values = (0 + 50)/2 = 25, but the most probable value of this random variable is 50, which will occur 90% of the time. To get the average probable value, we take the average of the values weighted by their probabilities. In this way, values with large probabilities count more than values with small probabilities. Formally, consider a discrete random variable X, which can take on values a1, a2, …, an, with probabilities p1, p2, …, pn, respectively. We define the expected value of X, which we write as E(X) (read “the expectation of X”) to be n

E ( X ) = ∑ ai pi = a1 p1 + a2 p2 + ... + an pn

(2.6)

i=1

Three points regarding E(X) are worth noting. First, the series n

∑ ai pi i=1

can have either a finite or an infinite number of terms, depending on the range of values that X can take on. Second, unlike an arithmetic average, the sum is not divided by the number of terms. Because probabilities are relative weights (probabilities are all scaled between 0 and 1), the division has already been done implicitly. And finally, except when all the pi are equal, this sum is not the same as the arithmetic average (see Chapter 3 for further discussion). For the three discrete random variables we have introduced—Bernoulli, binomial, and Poisson—the expected values are p, np, and λ, respectively. The Variance of a Discrete Random Variable

The expectation of a random variable describes the average, or central tendency, of the values. However, this number (like all averages) gives no insight into the spread, or variation, among the values. And, like the average family with 2.2 children, the expectation will not necessarily give an accurate depiction of an individual datum. In fact, for some discrete distributions such as the binomial or Poisson, none of the random variables may ever equal the expectation.

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For example, a binomial random variable that can take on the values –10 and +10, each value with probability 0.5, has the same expectation as a binomial random variable that can take on the values –1000 and +1000, each with probability = 0.5. Both have E(X) = 0, but in neither distribution will a random value of 0 ever be generated. Moreover, the observed values of the first distribution are much closer to the expected value than are the observed values of the second. Although the expectation accurately describes the midpoint of a distribution, we need some way to quantify the spread of the values around that midpoint. The variance of a random variable, which we write as σ2(X), is a measurement of how far the actual values of a random variable differ from the expected value. The variance of a random variable X is defined as

σ 2 ( X ) = E[X − E( X )]2 n ⎛ ⎞2 ⎜ = ∑ pi ⎜ai − ∑ ai pi ⎟⎟ ⎝ ⎠ i=1 i=1 n

(2.7)

As in Equation 2.6, E(X) is the expected value of X, and the ai’s are the different possible values of the variable X, each of which occurs with probability pi. To calculate σ2(X), we first calculate E(X), subtract this value from X, then square this difference. This is a basic measure of how much each value X differs from the expectation E(X).8 Because there are many possible values of X for random variables (e.g., two possible values for a Bernoulli random variable, n possible trial values for a binomial random variable, and infinitely many for a Poisson random variable), we repeat this subtraction and squaring for each possible value of X. Finally, we calculate the expectation of these squared deviates by following the same procedure as in Equation 2.6: each squared deviate is weighted by its probability of occurrence (pi) and then summed. Thus, in our example above, if Y is the binomial random variable that can take on values –10 and +10, each with P(Y) = 0.5, then σ2(Y) = 0.5(–10 – 0)2 + 0.5(10 – 0)2 = 100. Similarly, if Z is the binomial random variable that can take 8

You may be wondering why we bother to square the deviation X – E(X) once it is calculated. If you simply add up the unsquared deviations you will discover that the sum is always 0 because E(X) sits at the midpoint of all the values of X. We are interested in the magnitude of the deviation, rather than its sign, so we could use the sum of absolute values of the deviations ⌺|X – E(X)|. However, the algebra of absolute values is not as simple as that of squared terms, which also have better mathematical properties. The sum of the squared deviations, ⌺([X – E(X)]2), forms the basis for analysis of variance (see Footnote 1 in Chapter 10).

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on values –1000 and +1000, then σ2(Z) = 100,000. This result supports our intuition that Z has greater “spread” than Y. Finally, notice that if all of the values of the random variable are identical [i.e., X = E(X)], then σ2(X) = 0, and there is no variation in the data. For the three discrete random variables we have introduced—Bernoulli, binomial, and Poisson—the variances are p(1 – p), np(1 – p), and λ, respectively (Table 2.3).The expectation and the variance are the two most important descriptors of a random variable or probability distribution; Chapter 3 will discuss other summary measures of random variables. For now, we turn our attention to continuous variables.

Continuous Random Variables Many ecological and environmental variables cannot be described by discrete variables. For example, wing lengths of birds or pesticide concentrations in fish tissues can take on any value (bounded by an interval with appropriate upper and lower limits), and the precision of the measured value is limited only by available instrumentation. When we work with discrete random variables,

TABLE 2.3 Three discrete statistical distributions Distribution Probability value

E(X)

s2(X)

Bernoulli

P(X) = p

p(1 – p)

Binomial

⎛ n⎞ P( X ) = ⎜ ⎟ p X (1 − p)n− X ⎝ X⎠

np(1 – p)

Poisson

P (x ) =

λx − λ e x!

Comments

Use for dichotomous outcomes Use for number of successes in n independent trials Use for independent rare events where λ is the rate at which events occur in time or space

Ecological example

To reproduce or not, that is the question Presence or absence of species

Distribution of rare species across a landscape

The probability value equation determines the probability of obtaining a particular value X for each distribution. The expectation E(X) of the distribution of values is estimated by the mean or average of a sample. The variance σ2(X) is a measure of the spread or deviation of the observations from E(X). These distributions are for discrete variables, which are measured as integers or counts.

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we are able to define the total sample space as a set of possible discrete outcomes. However, when we work with continuous variables, we cannot identify all the possible events or outcomes as there are infinitely many of them (often uncountably infinitely many; see Footnote 4 in Chapter 1). Similarly, because observations can take on any value within the defined interval, it is difficult to define the probability of obtaining a particular value. We illustrate these issues as we describe our first type of continuous random variable, the uniform random variable. Uniform Random Variables

Both of the problems mentioned above—defining the appropriate sample space and obtaining the probability of any given value—are solvable. First, we recognize that our sample space is no longer discrete, but continuous. In a continuous sample space, we no longer consider discrete outcomes (such as X = 2), but instead focus on events that occur within a given subinterval (such as 1.5 < X < 2.5). The probability that an event occurs within a subinterval can itself be treated as an event, and our rules of probability continue to hold for such events. We start with a theoretical example: the closed unit interval, which contains all numbers between 0 and 1, including the two endpoints 0 and 1, and which we write as [0,1]. In this closed-unit interval, suppose that the probability of an event X occurring between 0 and 1/4 = p1, and the probability of this same event occurring between 1/2 and 3/4 = p2. By the rule that the probability of the union of two independent events equals the sum of their probabilities (see Chapter 1), the probability that X occurs in either of these two intervals (0 to 1/4 or 1/2 to 3/4) = p1 + p2. The second important rule is that all the probabilities of an event X in a continuous sample space must sum to 1 (this is the First Axiom of Probability). Suppose that, within the closed unit interval, all possible outcomes have equal probability (imagine, for example, a die with an infinite number of sides). Although we cannot define precisely the probability of obtaining the value 0.1 on a roll of this infinite die, we could divide the interval into 10 half-open intervals of equal length {[0,1/10), [1/10, 2/10),…,[9/10,1]} and calculate the probability of any roll being within one of these subintervals. As you might guess, the probability of a roll being within any one of these ten subintervals would be 0.1, and the sum of all the probabilities in this set = 10 × 0.1 = 1.0. Figure 2.4 illustrates this principle. In this example, the interval ranges from 0 to 10 (on the x-axis). Draw a line L parallel to the x-axis with a y-intercept of 0.1. For any subinterval U on this interval, the probability of a single roll of the die falling in that interval can be found by finding the area of the rectangle bounding the subinterval. This rectangle has length equal to the size of the subin-

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P(X)

0.10

Subinterval U = [3,4]

0 0

Figure 2.4 A uniform distribution on the interval [0,10]. In a continuous uniform distribution, the probability of an event occurring in a particular subinterval depends on the relative area in the subinterval; it is the same regardless of where that subinterval occurs within the bounded limits of the distribution. For example, if the distribution is bounded between 0 and 10, the probability that an event occurs in the subinterval [3,4] is the relative area that is bounded by that subinterval, which in this case is 0.10. The probability is the same for any other subinterval of the same size, such as [1,2], or [4,5]. If the subinterval chosen is larger, the probability of an event occurring in that subinterval is proportionally larger. For example the probability of an event occurring in the subinterval [3,5] is 0.20 (since 2 interval units of 10 are traversed), and is 0.6 in the subinterval [2,8].

terval and height = 0.1. If we divide the interval into u equal subintervals, the sum of all the areas of these subintervals would equal the sum of the area of the whole interval: 10 (length) × 0.1 (height) = 1.0. This graph also illustrates that the probability of any particular outcome a within a continuous sample space is 0, because the subinterval that contains only a is infinitely small, and an infinitely small number divided by a larger number = 0. We can now define a uniform random variable X with respect to any particular interval I. The probability that this uniform random variable X occurs in any subinterval U equals the product U × I. In the example illustrated in Figure 2.4, we define the following function to describe this uniform random variable: ⎧1/10, 0 ≤ x ≤ 10 f (x ) = ⎨ ⎩0 otherwise This function f(x) is called the probability density function (PDF) for this uniform distribution. In general, the PDF of a continuous random variable is found

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CHAPTER 2 Random Variables and Probability Distributions

by assigning probabilities that a continuous random variable X occurs within an interval I. The probability of X occurring within the interval I equals the area of the region bounded by I on the x-axis and f (x) on the y-axis. By the rules of probability, the total area under the curve described by the PDF = 1. We can also define a cumulative distribution function (CDF) of a random variable X as the function F(y) = P(X < y). The relationship between the PDF and the CDF is as follows: If X is a random variable with PDF f (x), then the CDF F(y) = P(X < Y) is equal to the area under f (x) in the interval x < y The CDF represents the tail probability—that is, the probability that a random variable X is less than or equal to some value y, [P(X) < y]—and is the same as the familiar P-value that we will discuss in Chapter 4. Figure 2.5 illustrates the PDF and the CDF for a uniform random variable on the closed unit interval.

1.0 X ~ Uniform[0,1] Probability density or cumulative density

0.8 Cumulative density function (CDF)

0.6 0.4 0.2

Probability density function (PDF)

0.0 0.0

0.2

0.4

0.6

0.8

1.0

Figure 2.5 Probability density function and cumulative distribution function for the uniform distribution measured over the closed unit interval [0,1]. The probability density function (PDF) shows the probability P(X) for any value X. In this continuous distribution, the exact probability of value X is technically 0, because the area under the curve is zero when measured at a single point. However, the area under the curve of any measurable subinterval is just the proportion of the total area under the curve, which by definition equals 1.0. In the uniform distribution, the probability of an event in any subinterval is the same regardless of where the subinterval is located. The cumulative density function (CDF) for this same distribution illustrates the cumulative area under the curve for the subinterval that is bounded at the low end by 0 and at the high end by 1.0. Because this is a uniform distribution, these probabilities accumulate in a linear fashion. When the end of the interval 1.0 is reached, CDF = 1.0, because the entire area under the curve has been included.

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Continuous Random Variables

The Expected Value of a Continuous Random Variable

We introduced the expected value of a random variable in the context of a discrete distribution. For a discrete random variable, n

E ( X ) = ∑ ai pi i=1

This calculation makes sense because each value of ai of X has an associated probability pi. However, for continuous distributions such as the uniform distribution, the probability of any particular observation = 0, and so we use the probabilities of events occurring within subintervals of the sample space. We take the same approach to find the expected value of a continuous random variable. To find the expected value of a continuous random variable, we will use very small subintervals of the sample space, which we will denote as Δx. For a probability density function f (x), the product of f (xi) and Δx gives the probability of an event occurring in the subinterval Δx, written as P(X = xi) = f (xi) Δx. This probability is the same as pi in the discrete case. As in Figure 2.4, the product of f (xi) and Δx describes the area of a very narrow rectangle. In the discrete case, we found the expected value by summing the product of each xi by its associated probability pi. In the continuous case, we will also find the expected value of a continuous random variable by summing the products of each xi and its associated probability f (xi)Δx. Obviously, the value of this sum will depend on the size of the small subinterval Δx. But it turns out that if our PDF f (x) has “reasonable” mathematical properties, and if we let Δx get smaller and smaller, then the sum n

∑ x i f (x i )Δx i=1

will approach a unique, limiting value. This limiting value = E(X) for a continuous random variable.9 For a uniform random variable X, where f (x) is defined on the interval [a,b], and where a < b, E(X) = (b + a)/2 9

If you’ve studied calculus, you will recognize this approach. For a continuous random variable X, where f (x) is differentiable within the sample space, E ( X ) = ∫ xf (x )dx

The integral represents the sum of the product x × f(x), where x becomes infinitely small in the limit.

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CHAPTER 2 Random Variables and Probability Distributions

The variance of a uniform random variable is σ2 (X ) =

(b − a)2 12

Normal Random Variables

Perhaps the most familiar probability distribution is the “bell curve”—the normal (or Gaussian) probability distribution. This distribution forms the theoretical basis for linear regression and analysis of variance (see Chapters 9 and 10), and it fits many empirical datasets. We will introduce the normal distribution with an example from the spider family Linyphiidae. Members of the different linyphiid genera can be distinguished by the length of their tibial 12

8 Frequency

4 2 0 0.136

0.162 0.188

0.214 0.240 0.266 0.292 0.318 Length of tibial spine (mm)

0.344 0.370 0.396

Figure 2.6 Normal distribution of a set of morphological measurements. Each observation in this histogram represents one of 50 measurements of tibial spine length in a sample of spiders (raw data in Table 3.1). The observations are grouped in “bins” that span 0.026-mm intervals; the height of each bar is the frequency (the number of observations that fell in that bin). Superimposed on the histogram is the normal distribution, with a mean of 0.253 and a standard deviation of 0.0039. Although the histogram does not conform perfectly to the normal distribution, the overall fit of the data is very good: the histogram exhibits a single central peak, an approximately symmetric distribution, and a steady decrease in the frequency of very large and very small measurements in the tails of the distribution.

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Continuous Random Variables

spines. An arachnologist measured 50 such spines and obtained the distribution illustrated in Figure 2.6. The histogram of these measurements illustrates several features that are characteristic of the normal distribution. First, notice that most of the observations are clustered around a central, or average tibial spine length. However, there are long tails of the histogram that extend to both the left and the right of the center. In a true normal distribution (in which we measured an infinite number of unfortunate spiders), these tails extend indefinitely in both directions, although the probability density quickly becomes vanishingly small as we move away from the center. In real datasets, the tails do not extend forever because we always have a limited amount of data, and because most measured variables cannot take on negative values. Finally, notice that the distribution is approximately symmetric: the left- and right-hand sides of the histogram are almost mirror images of one another. If we consider spine length to be a random variable X, we can use the normal probability density function to approximate this distribution. The normal distribution is described by two parameters, which we will call μ and σ, and so f (x) = f (μ,σ). The exact form of this function is not important here,10 but it has the properties that E(X) = μ, σ2(X) = σ2, and the distribution is symmetrical around μ. E(X) is the expectation and represents the central tendency of the data; σ2(X) is the variance and represents the spread of the obser-

If you’re craving the details, the PDF for the normal distribution is

f (x ) =

1 s 2p

1⎛ X−m ⎞ − ⎜ 2 ⎝ s ⎟⎠

where π is 3.14159…, e is the base of the natural logarithm (2.71828…), and μ and σ are the parameters defining the distribution. You can see from this formula that the more distant X is from μ, the larger the negative exponent of e, and hence the smaller the calculated probability for X. The CDF of this distribution, X

F(X ) =

∫ f (x)dx

−∞

does not have an analytic solution. Most statistics texts provide look-up tables for the CDF of the normal distribution. It also can be approximated using numerical integration techniques in standard software packages such as MatLab or R.

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CHAPTER 2 Random Variables and Probability Distributions

vations around the expectation. A random variable X described by this distribution is called a normal random variable, or a Gaussian random variable,11 and is written as X ∼ N(μ,σ)

(2.8)

Many different normal distributions can be created by specifying different values for μ and σ. However, statisticians often use a standard normal distribution, where μ = 0 and σ = 1. The associated standard normal random variable is usually referred to simply as Z. E(Z) = 0, and σ2(Z) = 1. Useful Properties of the Normal Distribution

The normal distribution has three useful properties. First, normal distributions can be added. If you have two independent normal random variables X and Y, their sum also is a normal random variable with E(X + Y) = E(X) + E(Y) and σ2(X + Y) = σ2(X) + σ2(Y). Second, normal distributions can be easily transformed with shift and change of scale operations. Consider two random variables X and Y. Let X ∼ N(μ,σ), and let Y = aX + b, where a and b are constants. We refer to the operation of

The Gaussian distribution is named for Karl Friedrich Gauss (1777–1855), one of the most important mathematicians in history. A child prodigy, he is reputed to have corrected an arithmetic error in his father’s payroll calculations at the age of 3. Gauss also proved the Fundamental Theorem of Algebra (every polynomial has a root of the form a + bi, where i = 冑苶 –1); the Fundamental Theorem of Arithmetic (every natural number can be represented as a unique Karl Friedrich Gauss product of prime numbers); formalized number theory; and, in 1801, developed the method of fitting a line to a set of points using least squares (see Chapter 9). Unfortunately, Gauss did not publish his method of least squares, and it is generally credited to Legendre, who published it 10 years later. Gauss found that the distribution of errors in the lines fit by least squares approximated what we now call the normal distribution, which was introduced nearly 100 years earlier by the American mathematician Abraham de Moivre (1667–1754) in his book The Doctrine of Chances. Because Gauss was the more famous of the two (at the time, the United States was a mathematical backwater), the normal probability distribution was originally referred to as the Gaussian distribution. De Moivre himself identified the normal distribution through his studies of the binomial distribution described earlier in this chapter. However, the modern name “normal” was not given to this distribution until the end of the nineteenth century (by the mathematician Poincaré), when the statistician Karl Pearson (1857–1936) rediscovered de Moivre’s work and showed that his discovery of this distribution predated Gauss’.

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Continuous Random Variables

multiplying X by a constant a as a change of scale operation because one unit of X becomes a units of Y; hence Y increases as a function of a. In contrast, the addition of a constant b to X is referred to as a shift operation because we simply move (shift) our random variable over b units along the x-axis by adding b to X. Shift and change of scale operations are illustrated in Figure 2.7. If X is a normal random variable, then a new random variable Y, created by either a change of scale operation, a shift operation, or both on the normal random variable X is also a normal random variable. Conveniently, the expectation and variance of the new random variable is a simple function of the shift and scale constants. For two random variables X ~ N(μ,σ) and Y = aX + b, we calculate E(Y) = aμ + b and σ2(Y) = a2σ2. Notice that the expectation of the new

2.0 Original data: tibial spine lengths

Frequency

1.5

Shift operation: tibial spine lengths + 5

1.0

Scale operation: tibial spine lengths × 5

0.5

0.0 0

2 3 4 Spine length (mm)

Figure 2.7 Shift and scale operations on a normal distribution. The normal distribution has two convenient algebraic properties. The first is a shift operation: if a constant b is added to a set of measurements with a mean μ, the mean of the new distribution is shifted to μ + b, but the variance is unaffected. The black curve is the normal distribution fit to a set of 200 measurements of spider tibial spine length (see Figure 2.6). The gray curve shows the shifted normal distribution after a value of 5 was added to each one of the original observations. The mean has shifted 5 units to the right, but the variance is unaltered. In a scale operation (blue curve), multiplying each observation by a constant a causes the mean to be increased by a factor of a, but the variance to be increased by a factor of a2. This curve is the normal distribution fit to the data after they have been multiplied by 5. The mean is shifted to a value of 5 times the original, and the variance has increased by a factor of 52 = 25.

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CHAPTER 2 Random Variables and Probability Distributions

random variable E(Y) is created by directly applying the shift and scale operations to E(X). However, the variance of the new random variable σ2(Y) is changed only by the scale operation. Intuitively, you can see that if all of the elements of a dataset are shifted by adding a constant, the variance should not change because the relative spread of the data has not been affected (see Figure 2.7). However, if those data are multiplied by a constant a (change of scale), the variance will be increased by the quantity a2; because the relative distance of each element from the expectation is now increased by a factor of a (Figure 2.7), this quantity is squared in our variance calculation. A final (and convenient) property of the normal distribution is the special case of a change of scale and shift operation in which a = 1/σ and b = –1(μ/σ): For X ∼ N(μ, σ), Y = (1/σ)X – μ/σ = (X – μ)/σ gives E(Y) = 0 and σ2(Y) = 1 which is a standard normal random variable. This is an especially useful result, because it means that any normal random variable can be transformed into a standard normal random variable. Moreover, any operation that can be applied to a standard normal random variable will apply to any normal random variable after it has been appropriately scaled and shifted. Other Continuous Random Variables

There are many other continuous random variables and associated probability distributions that ecologists and statisticians use. Two important ones are lognormal random variables and exponential random variables (Figure 2.8). A lognormal random variable X is a random variable such that its natural logarithm, ln(X), is a normal random variable. Many key ecological characteristics of organisms, such as body mass, are log-normally distributed.12 Like the normal distribution, the log-normal is described by two parameters, μ and σ. The expected value of a log-normal random variable is 2μ+σ 2 E( X ) = e 2

The most familiar ecological example of a log-normal distribution is the distribution of relative abundances of species in a community. If you take a large, random sample of individuals from a community, sort them according to species, and construct a histogram of the frequency of species represented in different abundance classes, the data will often resemble a normal distribution when the abundance classes are plotted on a logarithmic scale (Preston 1948). What is the explanation for this pattern? On the one hand, many non-biological datasets (such as the distribution of economic wealth among countries, or the distribution of “survival times” of drinking glasses in a busy restaurant) also follow a log-normal distribution. Therefore, the pattern may reflect a generic statistical response of exponentially increasing populations (a loga-

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Continuous Random Variables

(A)

(B) X ~ Log-normal(1,1) 0.06

X ~ Exponential(2)

0.30 0.25

P(X)

0.20 0.04 0.15 0.10 0.02 0.05 0.00

0.00 30

130

120

Figure 2.8 Log-normal and exponential distributions fit certain kinds of ecological data, such as species abundance distributions and seed dispersal distances. (A) The log-normal distribution is described by two parameters, a mean and a variance, both set to 1.0 in this example. (B) The exponential distribution is described by a single parameter b, set to 2.0 in this example. See Table 2.4 for the equations used with the log-normal and exponential distributions. Both the log-normal and the exponential distributions are asymmetric, with long right-hand tails that skew the distribution to the right.

and the variance of a log-normal distribution is ⎡ μ +σ 2 ⎤2 ⎡ 2 ⎤ σ 2 (X ) = ⎢e 2 ⎥ × ⎢e σ − 1⎥ ⎦ ⎣⎢ ⎦⎥ ⎣ When plotted on a logarithmic scale, the log-normal distribution shows a characteristic bell-shaped curve. However, if these same data are back-transformed to their original values (by applying the transformation eX), the resulting disrithmic phenomenon) to many independent factors (May 1975). On the other hand, specific biological mechanisms may be at work, including patch dynamics (Ugland and Gray 1982) or hierarchical niche partitioning (Sugihara 1980). The study of lognormal distributions is also complicated by sampling problems. Because rare species in a community may be missed in small samples, the shape of the species-abundance distribution changes with the intensity of sampling (Wilson 1993). Moreover, even in well-sampled communities, the tail of the frequency histogram may not fit a true lognormal distribution very well (Preston 1981). This lack of fit may arise because a large sample of a community will usually contain mixtures of resident and transient species (Magurran and Henderson 2003).

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CHAPTER 2 Random Variables and Probability Distributions

TABLE 2.4 Four continuous distributions Distribution

Probability value

E(X)

s2(X)

Comments

Uniform

P(a < X < b) = 1.0

b+a 2

(b − a)2 12

σ2

Use for equiprobable outcomes over interval [a,b] Generates a symmetric “bell curve” for continuous data Log-transformed data of rightskewed data are often fit by a normal distribution Continuous distribution analog of Poisson

Normal

1 σ 2π

Log-normal

1 σX 2π

Exponential

1 ⎛ X −μ ⎞ − ⎜ 2 ⎝ σ ⎟⎠

1 ⎛ ln( X )−μ ⎞ − ⎜ ⎟ 2⎝ ⎠ σ

P(X) = βe–βX

2μ+σ 2 e 2

1/β

⎡ μ +σ2 ⎤ × ⎡e σ2 − 1⎤ ⎥⎦ ⎢⎣e 2 ⎥⎦ ⎢⎣

1/β2

Ecological example

Even distribution of resources

Distribution of tibial spine lengths or other continuous size variables Distribution of species abundance classes

Seed dispersal distance

The Probability value equation determines the probability of obtaining a particular value X for each distribution. The expectation E(X) of the distribution of values is estimated by the mean or average of a sample. The variance σ2(X) is a measure of the spread or deviation of the observations from E(X). These distributions are for variables measured on a continuous scale that can take on any real number value, although the exponential distribution is limited to positive values.

tribution is skewed, with a long probability tail extending to the right. See Chapter 3 for a further discussion of measures of skewness, and Chapter 8 for more examples of data transformations. Exponential random variables are related to Poisson random variables. Recall that Poisson random variables describe the number of rare occurrences (e.g., counts), such as the number of arrivals in a constant time interval, or the number of individuals occurring in a fixed area. The “spaces” between discrete Poisson random variables, such as the time or distance between Poisson events, can be described as continuous exponential random variables. The probability distribution function for an exponential random variable X has only one parameter, usually written as β, and has the form P(X) = βe –βX. The expected value of an exponential random variable = 1/β, and its variance = 1/β2.13 Table 2.4 summarizes the properties of these common continuous distributions.

13 It is easy to simulate an exponential random variable on your computer by taking advantage of the fact that if U is a uniform random variable defined on the closed unit interval [0,1], then –ln(U)/β is an exponential random variable with parameter β.

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The Central Limit Theorem

Other continuous random variables and their probability distribution functions are used extensively in statistical analyses. These include the Student-t, chisquare, F, gamma, inverse gamma, and beta. We will discuss these later in the book when we use them with particular analytical techniques.

The Central Limit Theorem The Central Limit Theorem is one of the cornerstones of probability and statistical analysis.14 Here is a brief description of the theorem. Let Sn be the sum or the average of any set of n independent, identically distributed random variables Xi: n

Sn = ∑ X i i=1

each of which has the same expected value μ and all of which have the same variance σ2. Then, Sn has the expected value of nμ and variance nσ2. If we standardize Sn by subtracting the expected value from each observation and dividing by the square root of the variance, n

Sstd =

Sn − nμ σ n

∑ X i − nμ i=1

σ n

then the distribution of a set of Sstd values approximates a standard normal variable. 14

The initial formulation of the Central Limit Theorem is due to Abraham De Moivre (biographical information in Footnote 11) and Pierre Laplace. In 1733, De Moivre proved his version of the Central Limit Theorem for a Bernoulli random variable. Pierre Laplace (1749–1827) extended De Moivre’s result to any binary random variable. Laplace is better remembered for his Mécanique céleste (Celestial Mechanics), which translated Newton’s system of geometrical studies of mechanics into a system based on calculus. Abraham De Moivre According to Boyer’s History of Mathematics (1968), after Napoleon had read Mécanique céleste, he asked Laplace why there was no mention of God in the work. Laplace is said to have responded that he had no need for that hypothesis. Napoleon later appointed Laplace to be Minister of the Interior, but eventually dismissed him with the comment that “he carried the spirit of the infinitely small into the management of affairs” (fide Boyer 1968). The Russian mathematician Pafnuty Chebyshev (1821–1884) proved the Central Limit Theorem for any random variable, but his complex Pierre Laplace proof is virtually unknown today. The modern, accessible proof of the Central Limit Theorem is due to Chebyshev’s students Andrei Markov (1856–1922) and Alexander Lyapounov (1857–1918).

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This is a powerful result. The Central Limit Theorem asserts that standardizing any random variable that itself is a sum or average of a set of independent random variables results in a new random variable that is “nearly the same as”15 a standard normal one. We already used this technique when we generated a standard normal random variable (Z) from a normal random variable (X). The beauty of the Central Limit Theorem is that it allows us to use statistical tools that require our sample observations to be drawn from a sample space that is normally distributed, even though the underlying data themselves may not be normally distributed. The only caveats are that the sample size must be “large enough,”16 and that the observations themselves must be independent and all drawn from a distribution with common expectation and variance. We will demonstrate the importance of the Central Limit Theorem when we discuss the different statistical techniques used by ecologists and environmental scientists in Chapters 9–12.

Summary Random variables take on a variety of measurements, but their distributions can be characterized by their expectation and variance. Discrete distributions such as the Bernoulli, binomial, and Poisson apply to data that are discrete counts, whereas continuous distributions such as the uniform, normal, and exponen-

More formally, the Central Limit Theorem states that for any standardized variable Yi =

Si − n μ

σ n the area under the standard normal probability distribution over the open interval (a,b) equals lim P(a < Yi < b)

i→∞

An important question for practicing ecologists (and statisticians) is how quickly the probability P(a < Yi < b) converges to the area under the standard normal probability distribution. Most ecologists (and statisticians) would say that the sample size i should be at least 10, but recent studies suggest that i must exceed 10,000 before the two converge to even the first two decimal places! Fortunately, most statistical tests are fairly robust to the assumption of normality, so we can make use of the Central Limit Theorem even though our standardized data may not exhibit a perfectly normal distribution. Hoffmann-Jørgensen (1994; see also Chapter 5) provides a thorough and technical review of the Central Limit Theorem.

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Summary

tial apply to data measured on a continuous scale. Regardless of the underlying distribution, the Central Limit Theorem asserts that the sums or averages of large, independent samples will follow a normal distribution if they are standardized. For a wide variety of data, including those collected most commonly by ecologists and environmental scientists, the Central Limit Theorem supports the use of statistical tests that assume normal distributions.

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CHAPTER 3

Summary Statistics: Measures of Location and Spread

Data are the essence of scientific investigations, but rarely do we report all the data that we collect. Rather, we summarize our data using summary statistics. Biologists and statisticians distinguish between two kinds of summary statistics: measures of location and measures of spread. Measures of location illustrate where the majority of data are found; these measures include means, medians, and modes. In contrast, measures of spread describe how variable the data are; these measures include the sample standard deviation, variance, and standard errors. In this chapter, we introduce the most common summary statistics and illustrate how they arise directly from the Law of Large Numbers, one of the most important theorems of probability. Henceforth, we will adopt standard statistical notation when describing random variables and statistical quantities or estimators. Random variables will be designated as Y, where each individual observation is indexed with a subscript, Yi . The subscript i indicates the ith observation. The size of the sample will be denoted by n, and so i can take on any integer value between 1 and n. The – arithmetic mean is written as Y . Unknown parameters (or population statistics) of distributions, such as expected values and variances, will be written with Greek letters (such as μ for the expected value, σ2 for the expected variance, σ for the expected standard deviation), whereas statistical estimators of those – parameters (based on real data) will be written with italic letters (such as Y for the arithmetic mean, s2 for the sample variance, and s for the sample standard deviation). Throughout this chapter, we use as our example the data illustrated in Figure 2.6, the simulated measurement of tibial spines of 50 linyphiid spiders. These data, sorted in ascending order, are illustrated in Table 3.1.

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CHAPTER 3 Summary Statistics: Measures of Location and Spread

TABLE 3.1 Ordered measurements of tibial spines of 50 linyphiid spiders (millimeters) 0.155 0.184 0.199 0.202 0.206

0.207 0.208 0.212 0.212 0.215

0.219 0.219 0.221 0.223 0.226

0.228 0.228 0.229 0.235 0.238

0.241 0.243 0.247 0.247 0.248

0.249 0.250 0.251 0.253 0.258

0.263 0.268 0.270 0.274 0.275

0.276 0.277 0.280 0.286 0.289

0.292 0.292 0.296 0.301 0.306

0.307 0.308 0.328 0.329 0.368

This simulated dataset is used throughout this chapter to illustrate measures of summary statistics and probability distributions. Although raw data of this sort form the basis for all of our calculations in statistics, the raw data are rarely published because they are too extensive and too difficult to comprehend. Summary statistics, if they are properly used, concisely communicate and summarize patterns in raw data without enumerating each individual observation.

Measures of Location The Arithmetic Mean

There are many ways to summarize a set of data. The most familiar is the average, or arithmetic mean of the observations. The arithmetic mean is calculated as the sum of the observations (Yi) divided by the number of observations – (n) and is denoted by Y : n

Yi ∑ i=1

(3.1)

– For the data in Table 3.1, Y = 0.253. Equation 3.1 looks similar to, but is not quite equivalent to, Equation 2.6, which was used in Chapter 2 to calculate the expected value of a discrete random variable: n

E(Y ) = ∑Yi pi i =1

where the Yi’s are the values that the random variable can have, and the pi’s are their probabilities. For a continuous variable in which each Yi occurs only once, with pi = 1/n, Equations 3.1 and 2.6 give identical results. For example, let Spine length be the set consisting of the 50 observations in Table 3.1: Spine length = {0.155, 0.184, …, 0.329, 0.368}. If each element (or event) in Spine length is independent of all others, then the probability pi of any of these 50 independent observations is 1/50. Using Equation 2.6, we can calculate the expected value of Spine length to be n

( ) ∑Y p

EY =

i i

i =1

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Measures of Location

where Yi is the ith element and pi = 1/50. This sum, n

( ) ∑Y × 501

EY =

i =1

is now equivalent to Equation 3.1, used to calculate the arithmetic mean of n observations of a random variable Y: n

∑ i =1

piYi =

∑ i =1

Yi ×

1 1 = 50 50

∑Yi i =1

To calculate this expected value of Spine length, we used the formula for the expected value of a discrete random variable (Equation 2.6). However, the data given in Table 3.1 represent observations of a continuous, normal random variable. All we know about the expected value of a normal random variable is that it has some underlying true value, which we denote as μ. Does our calculated value of the mean of Spine length have any relationship to the unknown value of μ? If three conditions are satisfied, the arithmetic mean of the observations in our sample is an unbiased estimator of μ. These three conditions are: 1. Observations are made on randomly selected individuals. 2. Observations in the sample are independent of each other. 3. Observations are drawn from a larger population that can be described by a normal random variable. – The fact that Y of a sample approximates μ of the population from which the sample was drawn is a special case of the second fundamental theorem of probability, the Law of Large Numbers.1 Here is a description of the Law of Large Numbers. Consider an infinite set of random samples of size n, drawn from a random variable Y. Thus, Y1 is a sample from Y with 1 datum, {y1}. Y2 is a sample of size 2, {y1, y2}, etc. The Law of Large Numbers establishes that, as the sample size n increases, the arithmetic

The modern (or “strong”) version of the Law of Large Numbers was proven by the Russian mathematician Andrei Kolmogorov (1903–1987), who also studied Markov processes such as those used in modern computational Bayesian analysis (see Chapter 5) and fluid mechanics.

Andrei Kolmogorov

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CHAPTER 3 Summary Statistics: Measures of Location and Spread

mean of Yi (Equation 3.1) approaches the expected value of Y, E(Y). In mathematical notation, we write ⎞ ⎛ n yi ⎟ ⎜ lim ⎜ i =1 = Yn ⎟ = E(Y ) n→∞ ⎜ n ⎟ ⎟ ⎜ ⎠ ⎝

∑

(3.2)

In words, we say that as n gets very large, the average of the Yi’s equals E(Y) (see Figure 3.1). In our example, the tibial spine lengths of all individuals of linyphiid spiders in a population can be described as a normal random variable with expected value = μ. We cannot measure all of these (infinitely many) spines, but we can measure a subset of them; Table 3.1 gives n = 50 of these measurements. If each spine measured is from a single individual spider, each spider chosen for measurement is chosen at random, and there is no bias in our measurements, then the expected value for each observation should be the same (because they come from the same infinitely large population of spiders). The Law of Large Numbers states that the average spine length of our 50 measurements approximates the expected value of the spine length in the entire population. Hence, we can estimate the unknown expected value μ with the average of our observations. As Figure 3.1 shows, the estimate of the true population mean is more reliable as we accumulate more data. Other Means

The arithmetic average is not the only measure of location of a set of data. In some cases, the arithmetic average will generate unexpected answers. For example, suppose a population of mule deer (Odocoileus hemionus) increases in size by 10% in one year and 20% in the next year. What is the average population growth rate each year?2 The answer is not 15%! You can see this discrepancy by working through some numbers. Suppose the initial population size is 1000 deer. After one year, the population size (N1) will be (1.10) × 1000 = 1100. After the second year, the population size (N2) will be (1.20) × 1100 = 1320. However, if the average growth rate were 15% per 2

In this analysis, we use the finite rate of increase, λ, as the parameter for population growth. λ is a multiplier that operates on the population size each year, such that Nt+1 = λNt. Thus, if the population increases by 10% every year, λ = 1.10, and if the population decreases by 5% every year, λ = 0.95. A closely related measure of population growth rate is the instantaneous rate of increase, r, whose units are individuals/(individuals × time). Mathematically, λ = er and r = ln(λ). See Gotelli (2008) for more details.

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0.5 0.4

0.3 0.2 0.1

Population mean (0.253) Sample mean (last is 0.256) Confidence interval (last is 0.243 to 0.269)

0 0

100

200 300 Sample size

400

500

Figure 3.1 Illustration of the Law of Large Numbers and the construction of confidence intervals using the spider tibial spine data of Table 3.1. The population mean (0.253) is indicated by the dotted line. The sample mean for samples of increasing size n is indicated by the central solid line and illustrates the Law of Large Numbers: as the sample size increases, the sample mean approaches the true population mean. The upper and lower solid lines illustrate 95% confidence intervals about the sample mean. The width of the confidence interval decreases with increasing sample size. 95% of confidence intervals constructed in this way should contain the true population mean. Notice, however, that there are samples (between the arrows) for which the confidence interval does not include the true population mean. Curve constructed using algorithms and R code based on Blume and Royall (2003).

year, the population size would be (1.15) × 1000 = 1150 after one year and then (1.15) × 1150 = 1322.50 after 2 years. These numbers are close, but not identical; after several more years, the results diverge substantially. In Chapter 2, we introduced the log-normal distribution: if Y is a random variable with a log-normal distribution, then the random variable Z = ln(Y) is a normal random variable. If we calculate the arithmetic mean of Z, THE GEOMETRIC MEAN

1 n ∑ Zi n i=1

(3.3)

what is this value expressed in units of Y? First, recognize that if Z = ln(Y), then Y = e Z, where e is the base of the natural logarithm and equals ~2.71828… . – – Thus, the value of Z in units of Y is e Z . This so-called back-transformed mean is called the geometric mean and is written as GMY.

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CHAPTER 3 Summary Statistics: Measures of Location and Spread

The simplest way to calculate the geometric mean is to take the antilog of the arithmetic mean:

GM Y

⎡1 n ⎤ ⎢ ln(Yi )⎥ n ⎢ ⎥⎦ = e ⎣ i =1

∑

(3.4)

A nice feature of logarithms is that the sum of the logarithms of a set of numbers equals the logarithm of their products: ln(Y1) + ln(Y2) + ... = ln(Y1Y2...Yn). So another way of calculating the geometric mean is to take the nth root of the product of the observations: GM Y = n Y1Y2 ...Yn

(3.5)

Just as we have a special symbol for adding up a series of numbers: n

∑Yi = Y1+Y2 + ... + Yn i=1

we also have a special symbol for multiplying a series of numbers: n

∏Yi = Y1 × Y2 × ... × Yn i=1

Thus, we could also write our formula for the geometric mean as n

GM Y = n ∏ Yi i=1

Let’s see if the geometric mean of the population growth rates does a better job of predicting average population growth rate than the arithmetic average does. First, if we express population growth rates as multipliers, the annual growth rates of 10% and 20% become 1.10 and 1.20, and the natural logarithms of these two values are ln(1.10) = 0.09531 and ln(1.20) = 0.18232. The arithmetic average of these two numbers is 0.138815. Back-calculating gives us a geometric mean of GMY = e0.138815 = 1.14891, which is slightly less than the arithmetic mean of 1.20. Now we can calculate population growth rate over two years using this geometric mean growth rate. In the first year, the population would grow to (1.14891)× (1000) = 1148.91, and in the second year to (1.14891) × (1148.91) = 1319.99. This is the same answer we got with 10% growth in the first year and 20% growth in the second year [(1.10)× (1000)× (1.20)] = 1320. The values would match perfectly if we had not rounded the calculated growth rate. Notice also that although population size is always an integer variable (0.91 deer can be seen only in a theoretical forest), we treat it as a continuous variable to illustrate these calculations.

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Measures of Location

Why does GMY give us the correct answer? The reason is that population growth is a multiplicative process. Note that ⎛ N 2 ⎞ ⎛ N1 ⎞ N 2 ⎛ N 2 ⎞ ⎛ N1 ⎞ =⎜ ⎟ ×⎜ ⎟ ≠ ⎜ ⎟ +⎜ ⎟ N 0 ⎝ N1 ⎠ ⎝ N 0 ⎠ ⎝ N1 ⎠ ⎝ N 0 ⎠ However, numbers that are multiplied together on an arithmetic scale can be added together on a logarithmic scale. Thus ⎡⎛ N ⎞ ⎛ N ⎞ ⎤ ⎛N ⎞ ⎛N ⎞ ln ⎢⎜ 2 ⎟ × ⎜ 1 ⎟ ⎥ = ln⎜ 2 ⎟ + ln⎜ 1 ⎟ ⎝ N1 ⎠ ⎝ N0 ⎠ ⎢⎣⎝ N 1 ⎠ ⎝ N 0 ⎠ ⎥⎦ A second kind of average can be calculated in a similar way, using the reciprocal transformation (1/Y). The reciprocal of the arithmetic mean of the reciprocals of a set of observations is called the harmonic mean:3 THE HARMONIC MEAN

HY =

1 1 1 n ∑ Yi

(3.6)

For the spine data in Table 3.1, GMY = 0.249 and HY = 0.246. Both of these means are smaller than the arithmetic mean (0.253); in general, these means are ordered – as Y > GMY > HY. However, if all the observations are equal (Y1 = Y2 = Y3 = … – Yn), all three of these means are identical as well (Y = GMY = HY).

The harmonic mean turns up in conservation biology and population genetics in the calculation of effective population size, which is the equivalent size of a population with completely random mating. If the effective population size is small (< 50), random changes in allele frequency due to genetic drift potentially are important. If population size changes from one year to the next, the harmonic mean gives the effective population size. For example, suppose a stable population of 100 sea otters passes through a severe bottleneck and is reduced to a population size of 12 for a single year. Thus, the population sizes are 100, 100, 12, 100, 100, 100, 100, 100, 100, and 100. The arithmetic mean of these numbers is 91.2, but the harmonic mean is only 57.6, an effective population size at which genetic drift could be important. Not only is the harmonic mean less than the arithmetic mean, the harmonic mean is especially sensitive to extreme values that are small. Incidentally, sea otters on the Pacific coast of North America did pass through a severe population bottleneck when they were overhunted in the eighteenth and nineteenth centuries. Although sea otter populations have recovered in size, they still exhibit low genetic diversity, a reflection of this past bottleneck (Larson et al. 2002). (Photograph by Warren Worthington, soundwaves.usgs.gov/2002/07/.)

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CHAPTER 3 Summary Statistics: Measures of Location and Spread

Other Measures of Location: The Median and the Mode

Ecologists and environmental scientists commonly use two other measures of location, the median and the mode, to summarize datasets. The median is defined as the value of a set of ordered observations that has an equal number of observations above and below it. In other words, the median divides a dataset into two halves with equal number of observations in each half. For an odd number of observations, the median is simply the central observation. Thus, if we considered only the first 49 observations in our spine-length data, the median would be the 25th observation (0.248). But with an even number of observations, the median is defined as the midpoint between the (n/2)th and [(n/2)+1]th observation. If we consider all 50 observations in Table 3.1, the median would be the average of the 25th and 26th observations, or 0.2485. Mode = 0.237

Median = 0.248

12 Mean = 0.253 10

8 Frequency

0 0.150

0.175 0.200 0.225 0.250 0.275 0.300 Length of tibial spine (mm)

0.325 0.350 0.375

Figure 3.2 Histogram of the tibial spine data from Table 3.1 (n = 50) illustrating the arithmetic mean, median, and mode. The mean is the expectation of the data, calculated as the average of the continuous measurements. The median is the midpoint of the ordered set of observations. Half of all observations are larger than the median and half are smaller. The mode is the most frequent observation.

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Measures of Location

The mode, on the other hand, is the value of the observations that occurs most frequently in the sample. The mode can be read easily off of a histogram of the data, as it is the peak of the histogram. Figure 3.2 illustrates the arithmetic mean, the median, and the mode in a histogram of the tibial spine data. When to Use Each Measure of Location

Why choose one measure of location over another? The arithmetic mean (Equation 3.1) is the most commonly used measure of location, in part because it is familiar. A more important justification is that the Central Limit Theorem (Chapter 2) shows that arithmetic means of large samples of random variables conform to a normal or Gaussian distribution, even if the underlying random variable does not. This property makes it easy to test hypotheses on arithmetic means. The geometric mean (Equations 3.4 and 3.5) is more appropriate for describing multiplicative processes such as population growth rates or abundance classes of species (before they are logarithmically transformed; see the discussion of the log-normal distribution in Chapter 2 and the discussion of data transformations in Chapter 8). The harmonic mean (Equation 3.6) turns up in calculations used by population geneticists and conservation biologists. The median or the mode better describe the location of the data when distributions of observations cannot be fit to a standard probability distribution, or when there are extreme observations. This is because the arithmetic, geometric, and harmonic means are very sensitive to extreme (large or small) observations, whereas the median and the mode tend to fall in the middle of the distribution regardless of its spread and shape. In symmetrical distributions such as the normal distribution, the arithmetic mean, median, and mode all are equal. But in asymmetrical distributions, such as that shown in Figure 3.2 (a relatively small random sample from an underlying normal distribution), the mean occurs toward the largest tail of the distribution, the mode occurs in the heaviest part of the distribution, and the median occurs between the two.4

People also use different measures of location to support different points of view. For example, the average household income in the United States is considerably higher than the more typical (or median) income. This is because income has a log-normal distribution, so that averages are weighted by the long right-hand tail of the curve, representing the ultrarich. Pay attention to whether the mean, median, or mode of a data set is reported, and be suspicious if it is reported without any measure of spread or variation.

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CHAPTER 3 Summary Statistics: Measures of Location and Spread

Measures of Spread It is never sufficient simply to state the mean or other measure of location. Because there is variation in nature, and because there is a limit to the precision with which we can make measurements, we must also quantify and publish the spread, or variability, of our observations.

The Variance and the Standard Deviation

We introduced the concept of variance in Chapter 2. For a random variable Y, the variance σ2(Y) is a measurement of how far the observations of this random variable differ from the expected value. The variance is defined as E[Y – E(Y)]2 where E(Y) is the expected value of Y. As with the mean, the true variance of a – population is an unknown quantity. Just as we calculated an estimate Y of the population mean μ using our data, we can calculate an estimate s2 of the population variance σ2 using our data: s2 =

1 (Y − Y )2 n∑ i

(3.7)

This value is also referred to as the mean square. This term, along with its companion, the sum of squares, n

SSY = ∑ (Yi − Y )2

(3.8)

i=1

will crop up again when we discuss regression and analysis of variance in Chapters 9 and 10. And just as we defined the standard deviation σ of a random variable as the (positive) square root of its variance, we can estimate it as s =冑苳 s2. The square root transformation ensures that the units of standard deviation are the same as the units of the mean. – We noted earlier that the arithmetic mean Y provides an unbiased estimate of μ. By unbiased, we mean that if we sampled the population repeatedly (infinitely many times) and computed the arithmetic mean of each sample (regardless of sample size), the grand average of this set of arithmetic means should equal μ. However, our initial estimates of variance and standard deviation are not unbiased estimators of σ2 and σ, respectively. In particular, Equation 3.7 consistently underestimates the actual variance of the population. The bias in Equation 3.7 can be illustrated with a simple thought experiment. Suppose you draw a single observation Y1 from a population and try to estimate μ and σ2(Y). Your estimate of μ is the average of your observations, which in this case is simply Y1 itself. However, if you estimate σ2(Y) using Equation 3.7, your answer will always equal 0.0 because your lone observation is the same as your

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Measures of Spread

estimate of the mean! The problem is that, with a sample size of 1, we have already used our data to estimate μ, and we effectively have no additional information to estimate σ2(Y). This leads directly to the concept of degrees of freedom. The degrees of freedom represent the number of independent pieces of information that we have in a dataset for estimating statistical parameters. In a dataset of sample size 1, we do not have enough independent observations that can be used to estimate the variance. The unbiased estimate of the variance, referred to as the sample variance, is calculated by dividing the sums of squares by (n – 1) instead of dividing by n. Hence, the unbiased estimate of the variance is s2 =

1 (Yi − Y )2 n −1 ∑

(3.9)

and the unbiased estimate of the standard deviation, referred to as the sample standard deviation,5 is s=

1 (Yi − Y )2 n −1 ∑

(3.10)

Equations 3.9 and 3.10 adjust for the degrees of freedom in the calculation of the sample variance and the standard deviation. These equations also illustrate that you need at least two observations to estimate the variance of a distribution. For the tibial spine data given in Table 3.1, s2 = 0.0017 and s = 0.0417. The Standard Error of the Mean

Another measure of spread, used frequently by ecologists and environmental scientists, is the standard error of the mean. This measure of spread is abbreviated as sY– and is calculated by dividing the sample standard deviation by the square root of the sample size: sY =

s n

(3.11)

This unbiased estimator of the standard deviation is itself unbiased only for relatively large sample sizes (n > 30). For smaller sample sizes, Equation 3.10 modestly tends to underestimate the population value of σ (Gurland and Tripathi 1971). Rohlf and Sokal (1995) provide a look-up table of correction factors by which s should be multiplied if n < 30. In practice, most biologists do not apply these corrections. As long as the sample sizes of the groups being compared are not greatly different, no serious harm is done by ignoring the correction to s.

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CHAPTER 3 Summary Statistics: Measures of Location and Spread

Figure 3.3 Bar chart showing the arith-

Sample standard deviation Standard error of the mean

0.25

Spine length (mm)

metic mean for the spine data in Table 3.1 (n = 50), along with error bars indicating the sample standard deviation (left bar) and standard error of the mean (right bar). Whereas the standard deviation measures the variability of the individual measurements about the mean, the standard error measures the variability of the estimate of the mean itself. The standard error equals the standard deviation divided by n , so it will always be smaller than the standard deviation, often considerably so. Figure legends and captions should always provide sample sizes and indicate clearly whether the standard deviation or the standard error has been used to construct error bars.

0.30

0.20 0.15

0.10

0.05

0.00

The Law of Large Numbers proves that for an infinitely large number of observations, ΣYi /n approximates the population mean μ, where Yn = {Yi} is a sample of size n from a random variable Y with expected value E(Y). Similarly, the variance of Yn = σ2/n. Because the standard deviation is simply the square root of the variance, the standard deviation of Yn is σ2 σ = n n which is the same as the standard error of the mean. Therefore, the standard error of the mean is an estimate of the standard deviation of the population mean μ. Unfortunately, some scientists do not understand the distinction between the standard deviation (abbreviated in figure legends as SD) and the standard error of the mean (abbreviated as SE).6 Because the standard error of the mean is always smaller than the sample standard deviation, means reported with standard errors appear less variable than those reported with standard deviations (Figure 3.3). However, the decision as to whether to present the sample standard deviation s or the standard error of the mean sY– depends on what 6

You may have noticed that we referred to the standard error of the mean, and not simply the standard error. The standard error of the mean is equal to the standard deviation of a set of means. Similarly, we could compute the standard deviation of a set of variances or other summary statistics. Although it is uncommon to see other standard errors in ecological and environmental publications, there may be times when you need to report, or at least consider, other standard errors. In Figure 3.1, the standard error of the median = 1.2533 × sY–, and the standard error of the standard deviation = 0.7071× sY–. Sokal and Rohlf (1995) provide formulas for standard errors of other common statistics.

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inference you want the reader to draw. If your conclusions based on a single sample are representative of the entire population, then report the standard error of the mean. On the other hand, if the conclusions are limited to the sample at hand, it is more honest to report the sample standard deviation. Broad observational surveys covering large spatial scales with large number of samples more likely are representative of the entire population of interest (hence, report sY–), whereas small, controlled experiments with few replicates more likely are based on a unique (and possibly unrepresentative) group of individuals (hence, report s). We advocate the reporting of the sample standard deviation, s, which more accurately reflects the underlying variability of the actual data and makes fewer claims to generality. However, as long as you provide the sample size in your text, figure, or figure legend, readers can compute the standard error of the mean from the sample standard deviation and vice versa. Skewness, Kurtosis, and Central Moments

The standard deviation and the variance are special cases of what statisticians (and physicists) call central moments. A central moment (CM) is the average of the deviations of all observations in a dataset from the mean of the observations, raised to a power r: CM =

1 n (Y − Y )r n∑ i

(3.12)

i=1

In Equation 3.12, n is the number of observations, Yi is the value of each indi– vidual observation, Y is the arithmetic mean of the n observations, and r is a positive integer. The first central moment (r = 1) is the sum of the differences of each observation from the sample average (arithmetic mean), which always equals 0. The second central moment (r = 2) is the variance (Equation 3.5). The third central moment (r = 3) divided by the standard deviation cubed (s 3) is called the skewness (denoted as g1): g1 =

(Y − Y )3 3 ∑ i ns

(3.13)

i=1

Skewness describes how the sample differs in shape from a symmetrical distribution. A normal distribution has g1 = 0. A distribution for which g1 > 0 is rightskewed: there is a long tail of observations greater than (i.e., to the right of) the mean. In contrast, g1 < 0, is left-skewed: there is a long tail of observations less than (i.e., to the left of) the mean (Figure 3.4).

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CHAPTER 3 Summary Statistics: Measures of Location and Spread

Skewness = 6.9 0.06 Skewness = –0.05 P(X)

0.04

0.02

0.00 –20

–10

20 X

Figure 3.4 Continuous distributions illustrating skewness (g1). Skewness measures

the extent to which a distribution is asymmetric, with either a long right- or left-hand probability tail. The blue curve is the log-normal distribution illustrated in Figure 2.8; it has positive skewness, with many more observations to the right of the mean than to the left (a long right tail), and a skewness measure of 6.9. The black curve represents a sample of 1000 observations from a normal random variable with identical mean and standard deviation as the log-normal distribution. Because these data were drawn from a symmetric normal distribution, they have approximately the same number of observations on either side of the mean, and the measured skewness is approximately 0.

The kurtosis is based on the fourth central moment (r = 4): ⎡ 1 g2 = ⎢ 4 ⎢⎣ ns

⎤

i=1

⎦

∑ (Yi − Y )4 ⎥⎥ − 3

(3.14)

Kurtosis measures the extent to which a probability density is distributed in the tails versus the center of the distribution. Clumped or platykurtic distributions have g2 < 0; compared to a normal distribution, there is more probability mass in the center of the distribution, and less probability in the tails. In contrast, leptokurtic distributions have g2 > 0. Leptokurtic distributions have less probability mass in the center, and relatively fat probability tails (Figure 3.5). Although skewness and kurtosis were often reported in the ecological literature prior to the mid-1980s, it is uncommon to see these values reported now. Their statistical properties are not good: they are very sensitive to outliers, and

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Measures of Spread

0.04 Kurtosis = –0.11

P(X)

0.03

0.02

0.01 Kurtosis = 6.05 0.00 –8

–6

–4

–2

0 X

Figure 3.5 Distributions illustrating kurtosis (g2). Kurtosis measures the extent

to which the distribution is fat-tailed or thin-tailed compared to a standard normal distribution. Fat-tailed distributions are leptokurtic, and contain relative more area in the tails of the distribution and less in the center. Leptokurtic distributions have positive values for g2. Thin-tailed distributions are platykurtic, and contain relatively less area in the tails of the distribution and more in the center. Platykurtic distributions have negative values for g2. The black curve represents a sample of 1000 observations from a normal random variable with mean = 0 and standard deviation = 1 (X ~ N(0,1)); its kurtosis is nearly 0. The blue curve is a sample of 1000 observations from a t distribution with 3 degrees of freedom. The t distribution is leptokurtic and has a positive kurtosis (g2 = 6.05 in this example).

to differences in the mean of the distribution. Weiner and Solbrig (1984) discuss the problem of using measures of skewness in ecological studies. Quantiles

Another way to illustrate the spread of a distribution is to report its quantiles. We are all familiar with one kind of quantile, the percentile, because of its use in standardized testing. When a test score is reported as being in the 90th percentile, 90% of the scores are lower than the one being reported, and 10% are above it. Earlier in this chapter we saw another example of a percentile—the median, which is the value located at the 50th percentile of the data. In presentations of statistical data, we most commonly report upper and lower quartiles— the values for the 25th and 75th percentiles—and upper and lower deciles—the values for the 10th and 90th percentiles. These values for the spine data are illustrated concisely in a box plot (Figure 3.6). Unlike the variance and standard

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CHAPTER 3 Summary Statistics: Measures of Location and Spread

Figure 3.6 Box plot illustrating quantiles of data from Table 3.1 (n = 50). The line indicates the 50th percentile (median), and the box encompasses 50% of the data, from the 25th to the 75th percentile. The vertical lines extend from the 10th to the 90th percentile. Spine length (mm)

0.36

90th percentile (upper decile)

0.31

75th percentile (upper quartile)

0.26

50th percentile (median)

0.21

25th percentile (lower quartile)

0.16

10th percentile (lower decile)

0.11

deviation, the values of the quantiles do not depend on the values of the arithmetic mean or median. When distributions are asymmetric or contain outliers (extreme data points that are not characteristic of the distribution they were sampled from; see Chapter 8), box plots of quantiles can portray the distribution of the data more accurately than conventional plots of means and standard deviations. Using Measures of Spread

By themselves, measures of spread are not especially informative. Their primary utility is for comparing data from different populations or from different treatments within experiments. For example, analysis of variance (Chapter 10) uses the values of the sample variances to test hypotheses that experimental treatments differ from one another. The familiar t-test uses sample standard deviations to test hypotheses that the means of two populations differ from each other. It is not straightforward to compare variability itself across populations or treatment groups because the variance and standard deviation depend on the sample mean. However, by discounting the standard deviation by the mean, we can calculate an independent measure of variability, called the coefficient of variation, or CV. – The CV is simply the sample standard deviation divided by the mean, s/Y , and is conventionally multiplied by 100 to give a percentage. The CV for our spine data = 16.5%. If another population of spiders had a CV of tibial spine

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Some Philosophical Issues Surrounding Summary Statistics

length = 25%, we would say that our first population is somewhat less variable than the second population. A related index is the coefficient of dispersion, which is calculated as the – sample variance divided by the mean (s2/Y ). The coefficient of dispersion can be used with discrete data to assess whether individuals are clumped or hyperdispersed in space, or whether they are spaced at random as predicted by a Poisson distribution. Biological forces that violate independence will cause observed distributions to differ from those predicted by the Poisson. For example, some marine invertebrate larvae exhibit an aggregated settling response: once a juvenile occupies a patch, that patch becomes very attractive as a settlement surface for subsequent larvae (Crisp 1979). Compared to a Poisson distribution, these aggregated or clumped distributions will tend to have too many samples with high numbers of occurrence, and too many samples with 0 occurrences. In contrast, many ant colonies exhibit strong territoriality and kill or drive off other ants that try to establish colonies within the territory (Levings and Traniello 1981). This segregative behavior also will push the distribution away from the Poisson. In this case the colonies will be hyperdispersed: there will be too few samples in the 0 frequency class and too few samples with high numbers of occurrence. Because the variance and mean of a Poisson random variable both equal λ, the coefficient of dispersion (CD) for a Poisson random variable = λ/λ = 1. On the other hand, if the data are clumped or aggregated, CD > 1.0, and if the data are hyperdispersed or segregated, CD < 1.0. However, the analysis of spatial pattern with Poisson distributions can become complicated because the results depend not only on the degree of clumping or segregation of the organisms, but also on the size, number, and placement of the sampling units. Hurlbert (1990) discusses some of the issues involved in fitting spatial data to a Poisson distribution.

Some Philosophical Issues Surrounding Summary Statistics The fundamental summary statistics—the sample mean, standard deviation, and variance—are estimates of the actual population-level parameters, μ, σ, and σ 2, which we obtain directly from our data. Because we can never sample the entire population, we are forced to estimate these unknown parameters by – Y, s, and s2. In doing so, we make a fundamental assumption: that there is a true fixed value for each of these parameters. The Law of Large Numbers proves that if we sampled our population infinitely many times, the average of the infinitely – many Y ’s that we calculated from our infinitely many samples would equal μ. The Law of Large Numbers forms the foundation for what has come to be known as parametric, frequentist, or asymptotic statistics. Parametric statis-

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CHAPTER 3 Summary Statistics: Measures of Location and Spread

tics are so called because the assumption is that the measured variable can be described by a random variable or probability distribution of known form with defined, fixed parameters. Frequentist or asymptotic statistics are so called because we assume that if the experiment were repeated infinitely many times, the most frequent estimates of the parameters would converge on (reach an asymptote at) their true values. But what if this fundamental assumption—that the underlying parameters have true, fixed values—is false? For example, if our samples were taken over a long period of time, there might be changes in spine length of spider tibias because of phenotypic plasticity in growth, or even evolutionary change due to natural selection. Or, perhaps our samples were taken over a short period of time, but each spider came from a distinct microhabitat, for which there was a unique expectation and variance of spider tibia length. In such a case, is there any real meaning to an estimate of a single value for the average length of a tibial spine in the spider population? Bayesian statistics begin with the fundamental assumption that population-level parameters such as μ, σ, and σ2 are themselves random variables. A Bayesian analysis produces estimates not only of the values of the parameters but also of the inherent variability in these parameters. The distinction between the frequentist and Bayesian approaches is far from trivial, and has resulted in many years of acrimonious debate, first among statisticians, and more recently among ecologists. As we will see in Chapter 5, Bayesian estimates of parameters as random variables often require complex computer calculations. In contrast, frequentist estimates of parameters as fixed values use simple formulas that we have outlined in this chapter. Because of the computational complexity of the Bayesian estimates, it was initially unclear whether the results of frequentist and Bayesian analyses would be quantitatively different. However, with fast computers, we are now able to carry out complex Bayesian analyses. Under certain conditions, the results of the two types of analyses are quantitatively similar. The decision of which type of analysis to use, therefore, should be based more on a philosophical standpoint than on a quantitative outcome (Ellison 2004). However, the interpretation of statistical results may be quite different from the Bayesian and frequentist perspectives. An example of such a difference is the construction and interpretation of confidence intervals for parameter estimates.

Confidence Intervals Scientists often use the sample standard deviation to construct a confidence interval around the mean (see Figure 3.1). For a normally distributed random

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Confidence Intervals

variable, approximately 67% of the observations occur within ±1 standard deviation of the mean, and approximately 96% of the observations occur within ±2 standard deviations of the mean.7 We use this observation to create a 95% confidence interval, which for large samples is the interval bounded by (Y − 1.96sY , Y + 1.96sY ). What does this interval represent? It means that the probability that the true population mean μ falls within the confidence interval = 0.95: P(Y − 1.96sY ≤ μ ≤ Y + 1.96sY ) = 0.95

(3.15)

Because our sample mean and sample standard error of the mean are derived from a single sample, this confidence interval will change if we sample the population again (although if our sampling is random and unbiased, it should not change by very much). Thus, this expression is asserting that the probability that the true population mean μ falls within a single calculated confidence interval = 0.95. By extension, if we were to repeatedly sample the population (keeping the sample size constant), 5% of the time we would expect that the true population mean μ would lie outside of this confidence interval. Interpreting a confidence interval is tricky. A common misinterpretation of the confidence interval is “There is a 95% chance that the true population mean μ occurs within this interval.” Wrong. The confidence interval either does or does not contain μ; unlike Schrödinger’s quantum cat (see Footnote 9 in Chapter 1), μ cannot be both in and out of the confidence interval simultaneously. What you can say is that 95% of the time, an interval calculated in this way will contain the fixed value of μ. Thus, if you carried out your sampling experiment 100 times, and created 100 such confidence intervals, approximately 95 of them would contain μ and 5 would not (see Figure 3.1 for an example of when a 95% confidence interval does not include the true population mean μ). Blume and Royall (2003) provide further examples and a more detailed pedagogical description.

Use the “two standard deviation rule” when you read the scientific literature, and get into the habit of quickly estimating rough confidence intervals for sample data. For example, suppose you read in a paper that average nitrogen content of a sample of plant tissues was 3.4% ± 0.2, where 0.2 is the sample standard deviation. Two standard deviations = 0.4, which is then added to and subtracted from the mean. Therefore, approximately 95% of the observations were between 3.0% and 3.8%. You can use this same trick when you examine bar graphs in which the standard deviation is plotted as an error bar. This is an excellent way to use summary statistics to spot check reported statistical differences among groups.

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CHAPTER 3 Summary Statistics: Measures of Location and Spread

This rather convoluted explanation is not satisfying, and it is not exactly what you would like to assert when you construct a confidence interval. Intuitively, you would like to be saying how confident you are that the mean is inside of your interval (i.e., you’re 95% sure that the mean is in the interval). A frequentist statistician, however, can’t assert that. If there is a fixed population mean μ, then it’s either inside the interval or not, and the probability statement (Equation 3.15) asserts how probable it is that this particular confidence interval includes μ. On the other hand, a Bayesian statistician turns this around. Because the confidence interval is fixed (by your sample data), a Bayesian statistician can calculate the probability that the population mean (itself a random variable) occurs within the confidence interval. Bayesian statisticians refer to these intervals as credibility intervals, in order to distinguish them from frequentist confidence intervals. See Chapter 5, Ellison (1996), and Ellison and Dennis (2010) for further details. Generalized Confidence Intervals

We can, of course, construct any percentile confidence interval, such as a 90% confidence interval or a 50% confidence interval. The general formula for an n% confidence interval is

(

P(Y − t a[n−1]sY ≤ m ≤ Y + t a[n−1]sY ) = 1 – a

)

(3.16)

where tα[n–1] is the critical value of a t-distribution with probability P = α, and sample size n. This probability expresses the percentage of the area under the two tails of the curve of a t-distribution (Figure 3.7). For a standard normal curve (a t-distribution with n = ∞), 95% of the area under the curve lies within ±1.96 standard deviations of the mean. Thus 5% of the area (P = 0.05) under the curve remains in the two tails beyond the points ±1.96. So what is this t-distribution? Recall from Chapter 2 that an arithmetic transformation of a normal random variable is itself a normal random variable. Con– sider the set of sample means {Y k} resulting from a set of replicate groups of measurements of a normal random variable with unknown mean μ. The devi– ation of the sample means from the population mean Y k – μ is also a normal – random variable. If this latter random variable (Y k – μ) is divided by the unknown population standard deviation σ, the result is a standard normal random variable (mean = 0, standard deviation = 1). However, we don’t know the population standard deviation σ, and must instead divide the deviations of each

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Confidence Intervals

0.30 0.25

P(X)

0.20 0.15 2.5% of the area under the curve (lower tail of the distribution)

0.10

2.5% of the area under the curve (upper tail of the distribution)

0.05

95% of the observations

0.00 –8

–6

–4

–2

0 t

Figure 3.7 t-distribution illustrating that 95% of the observations, or probability mass lies within ±1.96 standard deviations of the mean (mean = 0) percentiles. The two tails of the distribution each contain 2.5% of the observations or probability mass of the distribution. Their sum is 5% of the observations, and the probability P = 0.05 that an observation falls within these tails. This distribution is identical to the t-distribution illustrated in Figure 3.5.

mean by the estimate of the standard error of the mean of each sample (sY–k). The resulting t-distribution is similar, but not identical, to a standard normal distribution. This t-distribution is leptokurtic, with longer and heavier tails than a standard normal distribution.8 8 This result was first demonstrated by the statistician W. S. Gossett, who published it using the pseudonym “Student.” Gossett at the time was employed at the Guinness Brewery, which did not allow its employees to publish trade secrets; hence the need for a pseudonym. This modified standard normal distribution, which Gossett called a t-distribution, is also known as the Student’s distribution, or the Student’s t-distribution. As the number of samples increases, the t-distribution approaches the standard normal distribution in shape. The construction of the t-distribution requires the specification of the sample size n and is normally written as t[n]. For n = ∞, t[∞] ~ N(0,1). Because a t-distribution for small n is leptokurtic (see Figure 3.5), the width of a confidence interval constructed from it will shrink as sample size increases. For example, for n = 10, 95% of the area under the curve falls between ± 2.228. For n = 100, 95% of the area under the curve falls between ±1.990. The resulting confidence interval is 12% wider for n = 10 than for n = 100.

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CHAPTER 3 Summary Statistics: Measures of Location and Spread

Summary Summary statistics describe expected values and variability of a random sample of data. Measures of location include the median, mode, and several means. If the samples are random or independent, the arithmetic mean is an unbiased estimator of the expected value of a normal distribution. The geometric mean and harmonic mean are also used in special circ*mstances. Measures of spread include the variance, standard deviation, standard error, and quantiles. These measures describe the variation of the observations around the expected value. Measures of spread can also be used to construct confidence or credibility intervals. The interpretations of these intervals differ between frequentist and Bayesian statisticians.

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CHAPTER 4

Framing and Testing Hypotheses

Hypotheses are potential explanations that can account for our observations of the external world. They usually describe cause-and-effect relationships between a proposed mechanism or process (the cause) and our observations (the effect). Observations are data—what we see or measure in the real world. Our goal in undertaking a scientific study is to understand the cause(s) of observable phenomena. Collecting observations is a means to that end: we accumulate different kinds of observations and use them to distinguish among different possible causes. Some scientists and statisticians distinguish between observations made during manipulative experiments and those made during observational or correlative studies. However, in most cases, the statistical treatment of such data is identical. The distinction lies in the confidence we can place in inferences1 drawn from those studies. Well-designed manipulative experiments allow us to be confident in our inferences; we have less confidence in data from poorly designed experiments, or from studies in which we were unable to directly manipulate variables. If observations are the “what” of science, hypotheses are the “how.” Whereas observations are taken from the real world, hypotheses need not be. Although our observations may suggest hypotheses, hypotheses can also come from the existing body of scientific literature, from the predictions of theoretical models, and from our own intuition and reasoning. However, not all descriptions of cause-and-effect relationships constitute valid scientific hypotheses. A scientific hypothesis must be testable: in other words, there should be some set of additional observations or experimental results that we could collect that would cause 1

In logic, an inference is a conclusion that is derived from premises. Scientists make inferences (draw conclusions) about causes based on their data. These conclusions may be suggested, or implied, by the data. But remember that it is the scientist who infers, and the data that imply.

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CHAPTER 4 Framing and Testing Hypotheses

us to modify, reject, or discard our working hypothesis.2 Metaphysical hypotheses, including the activities of omnipotent gods, do not qualify as scientific hypotheses because these explanations are taken on faith, and there are no observations that would cause a believer to reject these hypotheses.3 In addition to being testable, a good scientific hypothesis should generate novel predictions. These predictions can then be tested by collecting additional observations. However, the same set of observations may be predicted by more than one hypothesis. Although hypotheses are chosen to account for our initial observations, a good scientific hypothesis also should provide a unique set of predictions that do not emerge from other explanations. By focusing on these unique predictions, we can collect more quickly the critical data that will discriminate among the alternatives.

Scientific Methods The “scientific method” is the technique used to decide among hypotheses on the basis of observations and predictions. Most textbooks present only a single scientific method, but practicing scientists actually use several methods in their work. 2 A scientific hypothesis refers to a particular mechanism or cause-and-effect relationship; a scientific theory is much broader and more synthetic. In its early stages, not all elements of a scientific theory may be fully articulated, so that explicit hypotheses initially may not be possible. For example, Darwin’s theory of natural selection required a mechanism of inheritance that conserved traits from one generation to the next while still preserving variation among individuals. Darwin did not have an explanation for inheritance, and he discussed this weakness of his theory in The Origin of Species (1859). Darwin did not know that precisely such a mechanism had in fact been discovered by Gregor Mendel in his experimental studies (ca. 1856) of inheritance in pea plants. Ironically, Darwin’s grandfather, Erasmus Darwin, had published work on inheritance two generations earlier, in his Zoonomia, or the Laws of Organic Life (1794–1796). However, Erasmus Darwin used snapdragons as his experimental organism, whereas Mendel used pea plants. Inheritance of flower color is simpler in pea plants than in snapdragons, and Mendel was able to recognize the particulate nature of genes, whereas Erasmus Darwin could not. 3

Although many philosophies have attempted to bridge the gap between science and religion, the contradiction between reason and faith is a critical fault line separating the two. The early Christian philosopher Tertullian (∼155–222 AD) seized upon this contradiction and asserted “Credo quai absurdum est” (“I believe because it is absurd”). In Tertullian’s view, that the son of God died is to be believed because it is contradictory; and that he rose from the grave has certitude because it is impossible (Reese 1980).

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Scientific Methods

Deduction and Induction

Deduction and induction are two important modes of scientific reasoning, and both involve drawing inferences from data or models. Deduction proceeds from the general case to the specific case. The following set of statements provides an example of classic deduction: 1. All of the ants in the Harvard Forest belong to the genus Myrmica. 2. I sampled this particular ant in the Harvard Forest. 3. This particular ant is in the genus Myrmica. Statements 1 and 2 are usually referred to as the major and minor premises, and statement 3 is the conclusion. The set of three statements is called a syllogism, an important logical structure developed by Aristotle. Notice that the sequence of the syllogism proceeds from the general case (all of the ants in the Harvard Forest) to the specific case (the particular ant that was sampled). In contrast, induction proceeds from the specific case to the general case:4 1. All 25 of these ants are in the genus Myrmica. 2. All 25 of these ants were collected in the Harvard Forest. 3. All of the ants in the Harvard Forest are in the genus Myrmica.

The champion of the inductive method was Sir Francis Bacon (1561–1626), a major legal, philosophical, and political figure in Elizabethan England. He was a prominent member of parliament, and was knighted in 1603. Among scholars who question the authorship of Shakespeare’s works (the so-called anti-Stratfordians), some believe Bacon was the true author of Shakespeare’s plays, but the evidence isn’t very compelling. Bacon’s most imporSir Francis Bacon tant scientific writing is the Novum organum (1620), in which he urged the use of induction and empiricism as a way of knowing the world. This was an important philosophical break with the past, in which explorations of “natural philosophy” involved excessive reliance on deduction and on published authority (particularly the works of Aristotle). Bacon’s inductive method paved the way for the great scientific breakthroughs by Galileo and Newton in the Age of Reason. Near the end of his life, Bacon’s political fortunes took a turn for the worse; in 1621 he was convicted of accepting bribes and was removed from office. Bacon’s devotion to empiricism eventually did him in. Attempting to test the hypothesis that freezing slows the putrefaction of flesh, Bacon ventured out in the cold during the winter of 1626 to stuff a chicken with snow. He became badly chilled and died a few days later at the age of 65.

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CHAPTER 4 Framing and Testing Hypotheses

Some philosophers define deduction as certain inference and induction as probable inference. These definitions certainly fit our example of ants collected in the Harvard Forest. In the first set of statements (deduction), the conclusion must be logically true if the first two premises are true. But in the second case (induction), although the conclusion is likely to be true, it may be false; our confidence will increase with the size of our sample, as is always the case in statistical inference. Statistics, by its very nature, is an inductive process: we are always trying to draw general conclusions based on a specific, limited sample. Both induction and deduction are used in all models of scientific reasoning, but they receive different emphases. Even using the inductive method, we probably will use deduction to derive specific predictions from the general hypothesis in each turn of the cycle. Any scientific inquiry begins with an observation that we are trying to explain. The inductive method takes this observation and develops a single hypothesis to explain it. Bacon himself emphasized the importance of using the data to suggest the hypothesis, rather than relying on conventional wisdom, accepted authority, or abstract philosophical theory. Once the hypothesis is formulated, it generates—through deduction—further predictions. These predictions are then tested by collecting additional observations. If the new observations match the predictions, the hypothesis is supported. If not, the hypothesis must be modified to take into account both the original observation and the new observations. This cycle of hypothesis–prediction–observation is repeatedly traversed. After each cycle, the modified hypothesis should come closer to the truth5 (Figure 4.1). Two advantages of the inductive method are (1) it emphasizes the close link between data and theory; and (2) it explicitly builds and modifies hypotheses based on previous knowledge. The inductive method is confirmatory in that we

Ecologists and environmental scientists rely on induction when they use statistical software to fit non-linear (curvy) functions to data (see Chapter 9). The software requires that you specify not only the equation to be fit, but also a set of initial values for the unknown parameters. These initial values need to be “close to” the actual values because the algorithms are local estimators (i.e., they solve for local minima or maxima in the function). Thus, if the initial estimates are “far away” from the actual values of the function, the curve-fitting routines may either fail to converge on a solution, or will converge on a non-sensical one. Plotting the fitted curve along with the data is a good safeguard to confirm that the curve derived from the estimated parameters actually fits the original data.

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Scientific Methods

Initial observation

suggests

Prediction generates

Experiments, Data

New observations Hypothesis

NO (modify hypothesis) Do new observations match predictions?

YES (confirm hypothesis)

Figure 4.1 The inductive method. The cycle of hypothesis, prediction, and observation is repeatedly traversed. Hypothesis confirmation represents the theoretical endpoint of the process. Compare the inductive method to the hypothetico-deductive method (see Figure 4.4), in which multiple working hypotheses are proposed and emphasis is placed on falsification rather than verification.

seek data that support the hypothesis, and then we modify the hypothesis to conform with the accumulating data.6 There are also several disadvantages of the inductive method. Perhaps the most serious is that the inductive method considers only a single starting hypothesis; other hypotheses are only considered later, in response to additional data and observations. If we “get off on the wrong foot” and begin exploring an incorrect hypothesis, it may take a long time to arrive at the correct answer

6 The community ecologist Robert H. MacArthur (1930–1972) once wrote that the group of researchers interested in making ecology a science “arranges ecological data as examples testing the proposed theories and spends most of its time patching up the theories to account for as many of the data as possible” (MacArthur 1962). This quote characterizes much of the early theoretical work in community ecology. Later, theoretical ecology developed as a Robert H. MacArthur discipline in its own right, and some interesting lines of research blossomed without any reference to data or the real world. Ecologists disagree about whether such a large body of purely theoretical work has been good or bad for our science (Pielou 1981; Caswell 1988).

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“Accepted truth”

CHAPTER 4 Framing and Testing Hypotheses

through induction. In some cases we may never get there at all. In addition, the inductive method may encourage scientists to champion pet hypotheses and perhaps hang on to them long after they should have been discarded or radically modified (Loehle 1987). And finally, the inductive method—at least Bacon’s view of it—derives theory exclusively from empirical observations. However, many important theoretical insights have come from theoretical modeling, abstract reasoning, and plain old intuition. Important hypotheses in all sciences have often emerged well in advance of the critical data that are needed to test them.7 Modern-Day Induction: Bayesian Inference

The null hypothesis is the starting point of a scientific investigation. A null hypothesis tries to account for patterns in the data in the simplest way possible, which often means initially attributing variation in the data to randomness or measurement error. If that simple null hypothesis can be rejected, we can move on to entertain more complex hypotheses.8 Because the inductive method begins with an observation that suggests an hypothesis, how do we generate an appropriate null hypothesis? Bayesian inference represents a modern, updated version of the inductive method. The principals of Bayesian inference can be illustrated with a simple example. The photosynthetic response of leaves to increases in light intensity is a wellstudied problem. Imagine an experiment in which we grow 15 mangrove seedlings, each under a different light intensity (expressed as photosynthetic photon flux density, or PPFD, in μmol photons per m2 of leaf tissue exposed to 7 For example, in 1931 the Austrian physicist Wolfgang Pauli (1900–1958) hypothesized the existence of the neutrino, an electrically neutral particle with negligible mass, to account for apparent inconsistencies in the conservation of energy during radioactive decay. Empirical confirmation of the existence of neutrino did not come until 1956. 8

The preference for simple hypotheses over complex ones has a long history in science. Sir William of Ockham’s (1290–1349) Principle of Parsimony states that “[Entities] are not to be multiplied beyond necessity.” Ockham believed that unnecessarily complex hypotheses were vain and insulting to God. The Principle of Parsimony is sometimes known as Ockham’s Razor, the razor shearing away unnecessary complexity. Ockham lived an interesting life. He Sir William of Ockham was educated at Oxford and was a member of the Franciscan order. He was charged with heresy for some of the writing in his Master’s thesis. The charge was eventually dropped, but when Pope John XXII challenged the Franciscan doctrine of apostolic poverty, Ockham was excommunicated and fled to Bavaria. Ockham died in 1349, probably a victim of the bubonic plague epidemic.

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Scientific Methods

light each second) and measure the photosynthetic response of each plant (expressed as μmol CO2 fixed per m2 of leaf tissue exposed to light each second). We then plot the data with light intensity on the x-axis (the predictor variable) and photosynthetic rate on the y-axis (the response variable). Each point represents a different leaf for which we have recorded these two numbers. In the absence of any information about the relationship between light and photosynthetic rates, the simplest null hypothesis is that there is no relationship between these two variables (Figure 4.2). If we fit a line to this null hypothesis, the slope of the line would equal 0. If we collected data and found some other relationship between light availability and photosynthetic rate, we would then use those data to modify our hypothesis, following the inductive method. But is it really necessary to frame the null hypothesis as if you had no information at all? Using just a bit of knowledge about plant physiology, we can formulate a more realistic initial hypothesis. Specifically, we expect there to be some

Net assimilation rate –2 –1 (μmol CO2 m s )

6 5 4 3 2 1 0 0

500

1000 1500 Photosynthetically active radiation (μmol m–2 s–1)

2000

2500

Figure 4.2 Two null hypotheses for the relationship between light intensity (measured as photosynthetically active radiation) and photosynthetic rate (measured as net assimilation rate) in plants. The simplest null hypothesis is that there is no association between the two variables (dashed line). This null hypothesis is the starting point for a hypothetico-deductive approach that assumes no prior knowledge about the relationship between the variables and is the basis for a standard linear regression model (see Chapter 9). In contrast, the blue curve represents a Bayesian approach of bringing prior knowledge to create an informed null hypothesis. In this case, the “prior knowledge” is of plant physiology and photosynthesis. We expect that the assimilation rate will rise rapidly at first as light intensity increases, but then reach an asymptote or saturation level. Such a relationship can be described by a Michaelis-Menten equation [Y = kX/(D + X)], which includes parameters for an asymptotic assimilation rate (k) and a half saturation constant (D) that controls the steepness of the curve. Bayesian methods can incorporate this type of prior information into the analysis.

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CHAPTER 4 Framing and Testing Hypotheses

maximum photosynthetic rate that the plant can achieve. Beyond this point, increases in light intensity will not yield additional photosynthate, because some other factor, such as water or nutrients, becomes limiting. Even if these factors were supplied and the plant were grown under optimal conditions, photosynthetic rates will still level out because there are inherent limitations in the rates of biochemical processes and electron transfers that occur during photosynthesis. (In fact, if we keep increasing light intensity, excessive light energy can damage plant tissues and reduce photosynthesis. But in our example, we limit the upper range of light intensities to those that the plant can tolerate.) Thus, our informed null hypothesis is that the relationship between photosynthetic rate and light intensity should be non-linear, with an asymptote at high light intensities (see Figure 4.2). Real data could then be used to test the degree of support for this more realistic null hypothesis (Figure 4.3). To determine which null hypothesis to use, we also must ask what, precisely, is the point of the study? The simple null hypothesis (linear equation) is appropriate if we merely want to establish that a non-random relationship exists between light intensity and photosynthetic rate. The informed null hypothesis (Michaelis-Menten equation) is appropriate if we want to compare saturation curves among species or to test theoretical models that make quantitative predictions for the asymptote or half-saturation constant. Figures 4.2 and 4.3 illustrate how a modern-day inductivist, or Bayesian statistician, generates an hypothesis. The Bayesian approach is to use prior knowledge or information to generate and test hypotheses. In this example, the prior knowledge was derived from plant physiology and the expected shape of the light saturation curve. However, prior knowledge might also be based 7

Figure 4.3 Relationship between light intensity and photosynthetic rate. The data are measurements of net assimilation rate and photosynthetically active radiation for n = 15 young sun leaves of the mangrove Rhizophora mangle in Belize (Farnsworth and Ellison 1996b). A Michaelis-Menten equation of the form Y = kX/(D + X) was fit to the data. The parameter estimates ± 1 standard error are k = 7.3 ± 0.58 and D = 313 ± 86.6.

Net assimilation rate (μmol CO2 m–2 s–1)

6 5 4 3 2 1 0 0

500

1000

1500

Photosynthetically active radiation (μmol m–2 s–1)

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2000

Scientific Methods

on the extensive base of published literature on light saturation curves (Björkman 1981; Lambers et al. 1998). If we had empirical parameter estimates from other studies, we could quantify our prior estimates of the threshold and asymptote values for light saturation. These estimates could then be used to further specify the initial hypothesis for fitting the asymptote value to our experimental data. Use of prior knowledge in this way is different from Bacon’s view of induction, which was based entirely on an individual’s own experience. In a Baconian universe, if you had never studied plants before, you would have no direct evidence on the relationship between light and photosynthetic rate, and you would begin with a null hypothesis such as the flat line shown in Figure 4.2. This is actually the starting point for the hypothetico-deductive method presented in the next section. The strict Baconian interpretation of induction is the basis of the fundamental critique of the Bayesian approach: that the prior knowledge used to develop the initial model is arbitrary and subjective, and may be biased by preconceived notions of the investigator. Thus, the hypothetico-deductive method is viewed by some as more “objective” and hence more “scientific.” Bayesians counter this argument by asserting that the statistical null hypotheses and curve-fitting techniques used by hypothetico-deductivists are just as subjective; these methods only seem to be more objective because they are familiar and uncritically accepted. For a further discussion of these philosophical issues, see Ellison (1996, 2004), Dennis (1996), and Taper and Lele (2004). The Hypothetico-Deductive Method

The hypothetico-deductive method (Figure 4.4) developed from the works of Sir Isaac Newton and other seventeenth-century scientists and was championed by the philosopher of science Karl Popper.9 Like the inductive method, the hypothetico-deductive method begins with an initial observation that we are trying to explain. However, rather than positing a single hypothesis and working

Karl Popper

The Austrian philosopher of science Karl Popper (1902–1994) was the most articulate champion of the hypothetico-deductive method and falsifiability as the cornerstone of science. In The Logic of Scientific Discovery (1935), Popper argued that falsifiability is a more reliable criterion of truth than verifiability. In The Open Society and Its Enemies (1945), Popper defended democracy and criticized the totalitarian implications of induction and the political theories of Plato and Karl Marx.

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CHAPTER 4 Framing and Testing Hypotheses

Initial observation suggests Hypothesis A

Hypothesis B

Hypothesis C

Hypothesis D

Falsifiable predictions (unique to each hypothesis) Prediction A

Prediction B

Prediction C

Prediction D

New observations YES (repeat attempts to falsify) Do new observations match predictions? NO (falsify hypothesis) Hypothesis A

Hypothesis B

Hypothesis C

Multiple failed falsifications

Hypothesis D

“Accepted truth”

Figure 4.4 The hypothetico-deductive method. Multiple working hypotheses are proposed and their predictions tested with the goal of falsifying the incorrect hypotheses. The correct explanation is the one that stands up to repeated testing but fails to be falsified.

forward, the hypothetico-deductive method asks us to propose multiple, working hypotheses. All of these hypotheses account for the initial observation, but they each make additional unique predictions that can be tested by further experiments or observations. The goal of these tests is not to confirm, but to falsify, the hypotheses. Falsification eliminates some of the explanations, and the list is winnowed down to a smaller number of contenders. The cycle of predictions

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Scientific Methods

and new observations is repeated. However, the hypothetico-deductive method never confirms an hypothesis; the accepted scientific explanation is the hypothesis that successfully withstands repeated attempts to falsify it. The two advantages of the hypothetico-deductive method are: (1) it forces a consideration of multiple working hypotheses right from the start; and (2) it highlights the key predictive differences between them. In contrast to the inductive method, hypotheses do not have to be built up from the data, but can be developed independently or in parallel with data collection. The emphasis on falsification tends to produce simple, testable hypotheses, so that parsimonious explanations are considered first, and more complicated mechanisms only later.10 The disadvantages of the hypothetico-deductive method are that multiple working hypotheses may not always be available, particularly in the early stages of investigation. Even if multiple hypotheses are available, the method does not really work unless the “correct” hypothesis is among the alternatives. In contrast, the inductive method may begin with an incorrect hypothesis, but can reach the correct explanation through repeated modification of the original hypothesis, as informed by data collection. Another useful distinction is that the inductive method gains strength by comparing many datasets to a single hypothesis, whereas the hypothetico-deductive method is best for comparing a single dataset to multiple hypotheses. Finally, both the inductive method and hypothetico-deductive method place emphasis on a single correct hypothesis, making it difficult to evaluate cases in which multiple factors are at work. This is less of a problem with the inductive approach, because multiple explanations can be incorporated into more complex hypotheses.

The logic tree is a well-known variant of the hypothetico-deductive method that you may be familiar with from chemistry courses. The logic tree is a dichotomous decision tree in which different branches are followed depending on the results of experiments at each fork in the tree. The terminal branch tips of the tree represent the different hypotheses that are being tested. The logic tree also can be found in the familiar dichotomous taxonomic key for identifying to species unknown plants or animals: “If the animal has 3 pairs of walking legs, go to couplet x; if it has 4 or more pairs, go to couplet y.” The logic tree is not always practical for complex ecological hypotheses; there may be too many branch points, and they may not all be dichotomous. However, it is always an excellent exercise to try and place your ideas and experiments in such a comprehensive framework. Platt (1964) champions this method and points to its spectacular success in molecular biology; the discovery of the helical structure of DNA is a classic example of the hypothetico-deductive method (Watson and Crick 1953).

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CHAPTER 4 Framing and Testing Hypotheses

Neither scientific method is the correct one, and some philosophers of science deny that either scenario really describes how science operates.11 However, the hypothetico-deductive and inductive methods do characterize much science in the real world (as opposed to the abstract world of the philosophy of science). The reason for spending time on these models is to understand their relationship to statistical tests of an hypothesis.

Testing Statistical Hypotheses Statistical Hypotheses versus Scientific Hypotheses

Using statistics to test hypotheses is only a small facet of the scientific method, but it consumes a disproportionate amount of our time and journal space. We use statistics to describe patterns in our data, and then we use statistical tests to decide whether the predictions of an hypothesis are supported or not. Establishing hypotheses, articulating their predictions, designing and executing valid experiments, and collecting, organizing, and summarizing the data all occur before we use statistical tests. We emphasize that accepting or rejecting a statistical hypothesis is quite distinct from accepting or rejecting a scientific hypothesis. The statistical null hypothesis is usually one of “no pattern,” such as no difference between groups, or no relationship between two continuous variables. In contrast, the alternative hypothesis is that pattern exists. In other words, there

No discussion of Popper and the hypothetico-deductive method would be complete without mention of Popper’s philosophical nemesis, Thomas Kuhn (1922–1996). In The Structure of Scientific Revolutions (1962), Kuhn called into question the entire framework of hypothesis testing, and argued that it did not represent the way that science was done. Kuhn believed that science was done within the context of major paradigms, or research Thomas Kuhn frameworks, and that the domain of these paradigms was implicitly adopted by each generation of scientists. The “puzzle-solving” activities of scientists constitute “ordinary science,” in which empirical anomalies are reconciled with the existing paradigm. However, no paradigm can encompass all observations, and as anomalies accumulate, the paradigm becomes unwieldy. Eventually it collapses, and there is a scientific revolution in which an entirely new paradigm replaces the existing framework. Taking somewhat of an intermediate stance between Popper and Kuhn, the philosopher Imre Lakatos (1922–1974) thought that scientific research programs (SRPs) consisted of a core of central principles that generated a belt of surrounding hypotheses that make more specific predictions. The predictions of the hypotheses can be tested by the scientific method, but the core is not directly accessible (Lakatos 1978). Exchanges between Kuhn, Popper, Lakatos, and other philosophers of science can be read in Lakatos and Musgrave (1970). See also Horn (1986) for further discussion of these ideas.

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Testing Statistical Hypotheses

are distinct differences in measured values between groups, or a clear relationship exists between two continuous variables. You must ask how such patterns relate to the scientific hypothesis you are testing. For example, suppose you are evaluating the scientific hypothesis that waves scouring a rocky coast create empty space by removing competitively dominant invertebrate species. The open space can be colonized by competitively subordinate species that would otherwise be excluded. This hypothesis predicts that species diversity of marine invertebrates will change as a function of level of disturbance (Sousa 1979). You collect data on the number of species on disturbed and undisturbed rock surfaces. Using an appropriate statistical test, you find no difference in species richness in these two groups. In this case, you have failed to reject the statistical null hypothesis, and the pattern of the data fail to support one of the predictions of the disturbance hypothesis. Note, however, that absence of evidence is not evidence of absence; failure to reject a null hypothesis is not equivalent to accepting a null hypothesis (although it is often treated that way). Here is a second example in which the statistical pattern is the same, but the scientific conclusion is different. The ideal free distribution is an hypothesis that predicts that organisms move between habitats and adjust their density so that they have the same mean fitness in different habitats (Fretwell and Lucas 1970). One testable prediction of this hypothesis is that the fitness of organisms in different habitats is similar, even though population density may differ. Suppose you measure population growth rate of birds (an important component of avian fitness) in forest and field habitats as a test of this prediction (Gill et al. 2001). As in the first example, you fail to reject the statistical null hypothesis, so that there is no evidence that growth rates differ among habitats. But in this case, failing to reject the statistical null hypothesis actually supports a prediction of the ideal free distribution. Naturally, there are many additional observations and tests we would want to make to evaluate the disturbance hypothesis or the ideal free distribution. The point here is that the scientific and statistical hypotheses are distinct entities. In any study, you must determine whether supporting or refuting the statistical null hypothesis provides positive or negative evidence for the scientific hypothesis. Such a determination also influences profoundly how you set up your experimental study or observational sampling protocols. The distinction between the statistical null hypothesis and the scientific hypothesis is so important that we will return to it later in this chapter. Statistical Significance and P-Values

It is nearly universal to report the results of a statistical test in order to assert the importance of patterns we observe in the data we collect. A typical assertion is: “The control and treatment groups differed significantly from one another

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(P = 0.01).” What, precisely, does “P = 0.01” mean, and how does it relate to the concepts of probability that we introduced in Chapters 1 and 2? AN HYPOTHETICAL EXAMPLE: COMPARING MEANS A common assessment problem in environmental science is to determine whether or not human activities result in increased stress in animals. In vertebrates, stress can be measured as levels of the glucocorticoid hormones (GC) in the bloodstream or feces. For example, wolves that are not exposed to snowmobiles have 872.0 ng GC/g, whereas wolves exposed to snowmobiles have 1468.0 ng GC/g (Creel et al. 2002). Now, how do you decide whether this difference is large enough to be attributed to the presence of snowmobiles?12 Here is where you could conduct a conventional statistical test. Such tests can be very simple (such as the familiar t-test), or rather complex (such as tests for interaction terms in an analysis of variance). But all such statistical tests produce as their output a test statistic, which is just the numerical result of the test, and a probability value (or P-value) that is associated with the test statistic.

Before we can define the probability of a statistical test, we must first define the statistical null hypothesis, or H0. We noted above that scientists favor parsimonious or simple explanations over more complex ones. What is the simplest explanation to account for the difference in the means of the two groups? In our example, the simplest explanation is that the differences represent random variation between the groups and do not reflect any systematic effect of snowmobiles. In other words, if we were to divide the wolves into two groups but not expose individuals in either

THE STATISTICAL NULL HYPOTHESIS

Many people try to answer this question by simply comparing the means. However, we cannot evaluate a difference between means unless we also have some feeling for how much individuals within a treatment group differ. For example, if several of the individuals in the no-snowmobile group have GC levels as low as 200 ng/g and others have GC levels as high as 1544 ng/g (the average, remember, was 872), then a difference of 600 ng/g between the two exposure groups may not mean much. On the other hand, if most individuals in the no-snowmobile group have GC levels between 850 and 950 ng/g, then a 600 ng/g difference is substantial. As we discussed in Chapter 3, we need to know not only the difference in the means, but the variance about those means—the amount that a typical individual differs from its group mean. Without knowing something about the variance, we cannot say anything about whether differences between the means of two groups are meaningful.

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Testing Statistical Hypotheses

group to snowmobiles, we might still find that the means differ from each other. Remember that it is extremely unlikely that the means of two samples of numbers will be the same, even if they were sampled from a larger population using an identical process. Glucocorticoid levels will differ from one individual to another for many reasons that cannot be studied or controlled in this experiment, and all of this variation—including variation due to measurement error—is what we label random variation. We want to know if there is any evidence that the observed difference in the mean GC levels of the two groups is larger than we would expect given the random variation among individuals. Thus, a typical statistical null hypothesis is that “differences between groups are no greater than we would expect due to random variation.” We call this a statistical null hypothesis because the hypothesis is that a specific mechanism or force—some force other than random variation—does not operate. Once we state the statistical null hypothesis, we then define one or more alternatives to the null hypothesis. In our example, the natural alternative hypothesis is that the observed difference in the average GC levels of the two groups is too large to be accounted for by random variation among individuals. Notice that the alternative hypothesis is not that snowmobile exposure is responsible for an increase in GC! Instead, the alternative hypothesis is focused simply on the pattern that is present in the data. The investigator can infer mechanism from the pattern, but that inference is a separate step. The statistical test merely reveals whether the pattern is likely or unlikely, given that the null hypothesis is true. Our ability to assign causal mechanisms to those statistical patterns depends on the quality of our experimental design and our measurements. For example, suppose the group of wolves exposed to snowmobiles had also been hunted and chased by humans and their hunting dogs within the last day, whereas the unexposed group included wolves from a remote area uninhabited by humans. The statistical analysis would probably reveal significant differences in GC levels between the two groups regardless of exposure to snowmobiles. However, it would be dangerous to conclude that the difference between the means of the two groups was caused by snowmobiles, even though we can reject the statistical null hypothesis that the pattern is accounted for by random variation among individuals. In this case, the treatment effect is confounded with other differences between the control and treatment groups (exposure to hunting dogs) that are potentially related to stress levels. As we will discuss in THE ALTERNATIVE HYPOTHESIS

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CHAPTER 4 Framing and Testing Hypotheses

Chapters 6 and 7, an important goal of good experimental design is to avoid such confounded designs. If our experiment was designed and executed correctly, it may be safe to infer that the difference between the means is caused by the presence of snowmobiles. But even here, we cannot pin down the precise physiological mechanism if all we did was measure the GC levels of exposed and unexposed individuals. We would need much more detailed information on hormone physiology, blood chemistry, and the like if we want to get at the underlying mechanisms.13 Statistics help us establish convincing patterns, and from those patterns we can begin to draw inferences or conclusions about cause-and-effect relationships. In most tests, the alternative hypothesis is not explicitly stated because there is usually more than one alternative hypothesis that could account for the patterns in the data. Rather, we consider the set of alternatives to be “not H0.” In a Venn diagram, all outcomes of data can then be classified into either H0 or not H0. THE P-VALUE In many statistical analyses, we ask whether the null hypothesis of random variation among individuals can be rejected. The P-value is a guide to making that decision. A statistical P-value measures the probability that observed or more extreme differences would be found if the null hypothesis were true. Using the notation of conditional probability introduced in Chapter 1, P-value = P(data | H0). Suppose the P-value is relatively small (close to 0.0). Then it is unlikely (the probability is small) that the observed differences could have been obtained if the null hypothesis were true. In our example of wolves and snowmobiles, a low P-value would mean that it is unlikely that a difference of 600 ng/g in GC levels would have been observed between the exposed and unexposed groups if there was only random variation among individuals and no consistent effect of snowmobiles (i.e., if the null hypothesis is true). Therefore, with a small P-value, the results would be improbable given the null hypothesis, so we reject it. Because we had only one alternative hypothesis in our study, our conclusion is that snow-

13 Even if the physiological mechanisms were elucidated, there would still be questions about ultimate mechanisms at the molecular or genetic level. Whenever we propose a mechanism, there will always be lower-level processes that are not completely described by our explanation and have to be treated as a “black box.” However, not all higher-level processes can be explained successfully by reductionism to lower-level mechanisms.

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Testing Statistical Hypotheses

mobiles (or something associated with them) could be responsible for the difference between the treatment groups.14 On the other hand, suppose that the calculated P-value is relatively large (close to 1.0). Then it is likely that the observed difference could have occurred given 14

Accepting an alternative hypothesis based on this mechanism of testing a null hypothesis is an example of the fallacy of “affirming the consequent” (Barker 1989). Formally, the P-value = P(data | H0). If the null hypothesis is true, it would result in (or in the terms of logic, imply) a particular set of observations (here, the data). We can write this formally as H0 ⇒ null data, where the arrow is read as “implies.” If your observations are different from those expected under H0, then a low P-value suggests that H0 ⇒ your data, where the crossed arrow is read as “does not imply.” Because you have set up only one alternative hypothesis, Ha, then you are further asserting that Ha = ¬H0 (where the symbol ¬ means “not”), and the only possibilities for data are those data possible under H0 (“null data”) and those not possible under H0 (“¬null data” = “your data”). Thus, you are asserting the following logical progression: 1. 2. 3. 4.

Given: H0 ⇒ null data Observe: ¬null data Conclude: ¬null data ⇒ ¬H0 Thus: ¬H0 (= Ha) ⇒ ¬null data

But really, all you can conclude is point 3: ¬null data ⇒ ¬H0 (the so-called contrapositive of 1). In 3, the alternative hypothesis (Ha) is the “consequent,” and you cannot assert its truth simply by observing its “predicate” (¬null data in 3); many other possible causes could have yielded your results (¬null data). You can affirm the consequent (assert Ha is true) if and only if there is only one possible alternative to your null hypothesis. In the simplest case, where H0 asserts “no effect” and Ha asserts “some effect,” proceeding from 3 to 4 makes sense. But biologically, it is usually of more interest to know what is the actual effect (as opposed to simply showing there is “some effect”). Consider the ant example earlier in this chapter. Let H0 = all 25 ants in the Harvard Forest are Myrmica, and Ha = 10 ants in the forest are not Myrmica. If you collect a specimen of Camponotus in the forest, you can conclude that the data imply that the null hypothesis is false (observation of a Camponotus in the forest ⇒ ¬H0). But you cannot draw any conclusion about the alternative hypothesis. You could support a less stringent alternative hypothesis, Ha = not all ants in the forest are Myrmica, but affirming this alternative hypothesis does not tell you anything about the actual distribution of ants in the forest, or the identity of the species and genera that are present. This is more than splitting logical hairs. Many scientists appear to believe that when they report a P-value that they are giving the probability of observing the null hypothesis given the data [P(H0 | data)] or the probability that the alternative hypothesis is false, given the data [1 – P(Ha | data)]. But, in fact, they are reporting something completely different—the probability of observing the data given the null hypothesis: P(data | H0). Unfortunately, as we saw in Chapter 1, P(data | H0) ≠ P(H0 | data) ≠ 1 – P(Ha | data); in the words of the immortal anonymous philosopher from Maine, you can’t get there from here. However, it is possible to compute directly P(H0 | data) or P(Ha | data) using Bayes’ Theorem (see Chapter 1) and the Bayesian methods outlined in Chapter 5.

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that the null hypothesis is true. In this example, a large P-value would mean that a 600-ng/g difference in GC levels likely would have been observed between the exposed and unexposed groups even if snowmobiles had no effect and there was only random variation among individuals. That is, with a large P-value, the observed results would be likely under the null hypothesis, so we do not have sufficient evidence to reject it. Our conclusion is that differences in GC levels between the two groups can be most parsimoniously attributed to random variation among individuals. Keep in mind that when we calculate a statistical P-value, we are viewing the data through the lens of the null hypothesis. If the patterns in our data are likely under the null hypothesis (large P-value), we have no reason to reject the null hypothesis in favor of more complex explanations. On the other hand, if the patterns are unlikely under the null hypothesis (small P-value), it is more parsimonious to reject the null hypothesis and conclude that something more than random variation among subjects contributes to the results. WHAT DETERMINES THE P-VALUE?

The calculated P-value depends on three things: the number of observations in the samples (n), the difference between the means – – of the samples (Yi – Yj), and the level of variation among individuals (s2). The more observations in a sample, the lower the P-value, because the more data we have, the more likely it is we are estimating the true population means and can detect a real difference between them, if it exists (see the Law of Large Numbers in Chapter 3). The P-value also will be lower the more different the two groups are in the variable we are measuring. Thus, a 10-ng/g difference in mean GC levels between control and treatment groups will generate a lower P-value than a 2-ng/g difference, all other things being equal. Finally, the P-value will be lower if the variance among individuals within a treatment group is small. The less variation there is from one individual to the next, the easier it will be to detect differences among groups. In the extreme case, if the GC levels for all individuals within the group of wolves exposed to snowmobiles were identical, and the GC levels for all individuals within the unexposed group were identical, then any difference in the means of the two groups, no matter how small, would generate a low P-value. WHEN IS A P-VALUE SMALL ENOUGH? In our example, we obtained a P-value = 0.01 for the probability of obtaining the observed difference in GC levels between wolves exposed to and not exposed to snowmobiles. Thus, if the null hypothesis were true and there was only random variation among individuals in the data, the chance of finding a 600-ng/g difference in GC between exposed and unexposed groups is only 1 in 100. Stated another way, if the null hypothesis were

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Testing Statistical Hypotheses

true, and we conducted this experiment 100 times, using different subjects each time, in only one of the experiments would we expect to see a difference as large or larger than what we actually observed. Therefore, it seems unlikely the null hypothesis is true, and we reject it. If our experiment was properly designed, we can safely conclude that snowmobiles cause increases in GC levels, although we cannot specify what it is about snowmobiles that causes this response. On the other hand, if the calculated statistical probability were P = 0.88, then we would expect a result similar to what we found in 88 out of 100 experiments due to random variation among individuals; our observed result would not be at all unusual under the null hypothesis, and there would be no reason to reject it. But what is the precise cutoff point that we should use in making the decision to reject or not reject the null hypothesis? This is a judgment call, as there is no natural critical value below which we should always reject the null hypothesis and above which we should never reject it. However, after many decades of custom, tradition, and vigilant enforcement by editors and journal reviewers, the operational critical value for making these decisions equals 0.05. In other words, if the statistical probability P ≤ 0.05, the convention is to reject the null hypothesis, and if the statistical probability P > 0.05, the null hypothesis is not rejected. When scientists report that a particular result is “significant,” they mean that they rejected the null hypothesis with a P-value ≤ 0.05.15 A little reflection should convince you that a critical value of 0.05 is relatively low. If you used this rule in your everyday life, you would never take an umbrella with you unless the forecast for rain was at least 95%. You would get wet a lot more often than your friends and neighbors. On the other hand, if your friends and neighbors saw you carrying your umbrella, they could be pretty confident of rain. In other words, setting a critical value = 0.05 as the standard for rejecting a null hypothesis is very conservative. We require the evidence to be very strong in order to reject the statistical null hypothesis. Some investigators are unhappy about using an arbitrary critical value, and about setting it as low as 0.05. After all, most of us would take an umbrella with a 90% forecast of rain, so why shouldn’t we be a bit less rigid in our standard for rejecting the null hypothesis? Perhaps we should set the critical critical value = 0.10, or perhaps we should use different critical values for different kinds of data and questions. 15

When scientists discuss “significant” results in their work, they are really speaking about how confident they are that a statistical null hypothesis has been correctly rejected. But the public equates “significant” with “important.” This distinction causes no end of confusion, and it is one of the reasons that scientists have such a hard time communicating their ideas clearly in the popular press.

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A defense of the 0.05-cutoff is the observation that scientific standards need to be high so that investigators can build confidently on the work of others. If the null hypothesis is rejected with more liberal standards, there is a greater risk of falsely rejecting a true null hypothesis (a Type I error, described in more detail below). If we are trying to build hypotheses and scientific theories based on the data and results of others, such mistakes slow down scientific progress. By using a low critical value, we can be confident that the patterns in the data are quite strong. However, even a low critical value is not a safeguard against a poorly designed experiment or study. In such cases, the null hypothesis may be rejected, but the patterns in the data reflect flaws in the sampling or manipulations, not underlying biological differences that we are seeking to understand. Perhaps the strongest argument in favor of requiring a low critical value is that we humans are psychologically predisposed to recognizing and seeing patterns in our data, even when they don’t exist. Our vertebrate sensory system is adapted for organizing data and observations into “useful” patterns, generating a built-in bias toward rejecting null hypotheses and seeing patterns where there is really randomness (Sale 1984).16 A low critical value is a safeguard against such activity. A low critical value also helps act as a gatekeeper on the rate of scientific publications because non-significant results are much less likely to be reported or published.17 We emphasize, however, that no law requires a critical value to be ≤ 0.05 in order for the results to be declared significant. In many cases, it may be more useful to report the exact P-value and let the readers decide for themselves how important the results are. However, the practical reality is that reviewers and editors will usually not allow you to discuss mechanisms that are not supported by a P ≤ 0.05 result.

A fascinating illustration of this is to ask a friend to draw a set of 25 randomly located points on a piece of paper. If you compare the distribution of those points to a set of truly random points generated by a computer, you will often find that the drawings are distinctly non-random. People have a tendency to space the points too evenly across the paper, whereas a truly random pattern generates apparent “clumps” and “holes.” Given this tendency to see patterns everywhere, we should use a low critical value to ensure we are not deceiving ourselves. 17 The well-known tendency for journals to reject papers with non-significant results (Murtaugh 2002a) and authors to therefore not bother trying to publish them is not a good thing. In the hypothetico-deductive method, science progresses through the elimination of alternative hypotheses, and this can often be done when we fail to reject a null hypothesis. However, this approach requires authors to specify and test the unique predictions that are made by competing alternative hypotheses. Statistical tests based on H0 versus not H0 do not often allow for this kind of specificity.

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Testing Statistical Hypotheses

The biggest difficulty in using P-values results from the failure to distinguish statistical null hypotheses from scientific hypotheses. Remember that a scientific hypothesis poses a formal mechanism to account for patterns in the data. In this case, our scientific hypothesis is that snowmobiles cause stress in wolves, which we propose to test by measuring GC levels. Higher levels of GC might come about by complex changes in physiology that lead to changes in GC production when an animal is under stress. In contrast, the statistical null hypothesis is a statement about patterns in the data and the likelihood that these patterns could arise by chance or random processes that are not related to the factors we are explicitly studying. We use the methods of probability when deciding whether or not to reject the statistical null hypothesis; think of this process as a method for establishing pattern in the data. Next, we draw a conclusion about the validity of our scientific hypothesis based on the statistical pattern in this data. The strength of this inference depends very much on the details of the experiment and sampling design. In a well-designed and replicated experiment that includes appropriate controls and in which individuals have been assigned randomly to clear-cut treatments, we can be fairly confident about our inferences and our ability to evaluate the scientific hypothesis we are considering. However, in a sampling study in which we have not manipulated any variables but have simply measured differences among groups, it is difficult to make solid inferences about the underlying scientific hypotheses, even if we have rejected the statistical null hypothesis.18 We think the more general issue is not the particular critical value that is chosen, but whether we always should be using an hypothesis-testing framework. Certainly, for many questions statistical hypothesis tests are a powerful way to establish what patterns do or do not exist in the data. But in many studies, the real issue may not be hypothesis testing, but parameter estimation. For example, in the stress study, it may be more important to determine the range of GC levels expected for wolves exposed to snowmobiles rather than merely to establish that snowmobiles significantly increases GC levels. We also should establish the level of confidence or certainty in our parameter estimates. STATISTICAL HYPOTHESES VERSUS SCIENTIFIC HYPOTHESES REDUX

18 In contrast to the example of the snowmobiles and wolves, suppose we measured the GC levels of 10 randomly chosen old wolves and 10 randomly chosen young ones. Could we be as confident about our inferences as in the snowmobile experiment? Why or why not? What are the differences, if any, between experiments in which we manipulate individuals in different groups (exposed wolves versus unexposed wolves) and sampling surveys in which we measure variation among groups but do not directly manipulate or change conditions for those groups (old wolves versus young wolves)?

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Errors in Hypothesis Testing

Although statistics involves many precise calculations, it is important not to lose sight of the fact that statistics is a discipline steeped in uncertainty. We are trying to use limited and incomplete data to make inferences about underlying mechanisms that we may understand only partially. In reality, the statistical null hypothesis is either true or false; if we had complete and perfect information, we would know whether or not it were true and we would not need statistics to tell us. Instead, we have only our data and methods of statistical inference to decide whether or not to reject the statistical null hypothesis. This leads to an interesting 2 × 2 table of possible outcomes whenever we test a statistical null hypothesis (Table 4.1) Ideally, we would like to end up in either the upper left or lower right cells of Table 4.1. In other words, when there is only random variation in our data, we would hope to not reject the statistical null hypothesis (upper left cell), and when there is something more, we would hope to reject it (lower right cell). However, we may find ourselves in one of the other two cells, which correspond to the two kinds of errors that can be made in a statistical decision. TYPE I ERROR If we falsely reject a null hypothesis that is true (upper right cell in Table 4.1), we have made a false claim that some factor above and beyond random variation is causing patterns in our data. This is a Type I error, and by convention, the probability of committing a Type I error is denoted by α. When you calculate a statistical P-value, you are actually estimating α. So, a more precise

TABLE 4.1 The quadripartite world of statistical testing H0 true H0 false

Retain H0

Reject H0

Correct decision Type II error (β)

Type I error (α) Correct decision

Underlying null hypotheses are either true or false, but in the real world we must use sampling and limited data to make a decision to accept or reject the null hypothesis. Whenever a statistical decision is made, one of four outcomes will result. A correct decision results when we retain a null hypothesis that is true (upper left-hand corner) or reject a null hypothesis that is false (lower right-hand corner). The other two possibilities represent errors in the decision process. If we reject a null hypothesis that is true, we have committed a Type I error (upper right-hand corner). Standard parametric tests seek to control α, the probability of a Type I error. If we retain a null hypothesis that is false, we have committed a Type II error (lower left-hand corner). The probability of a Type II error is β.

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Testing Statistical Hypotheses

definition of a P-value is that it is the chance we will make a Type I error by falsely rejecting a true null hypothesis.19 This definition lends further support for asserting statistical significance only when the P-value is very small. The smaller the P-value, the more confident we can be that we will not commit a Type I error if we reject H0. In the glucocorticoid example, the risk of making a Type I error by rejecting the null hypothesis is 1%. As we noted before, scientific publications use a standard of a maximum of a 5% risk of Type I error for rejecting a null hypothesis. In environmental impact assessment, a Type I error would be a “false positive” in which, for example, an effect of a pollutant on human health is reported but does not, in fact, exist. TYPE II ERROR AND STATISTICAL POWER The lower left cell in Table 4.1 represents a Type II error. In this case, the investigator has incorrectly failed to reject a null hypothesis that is false. In other words, there are systematic differences between the groups being compared, but the investigator has failed to reject the null hypothesis and has concluded incorrectly that only random variation among observations is present. By convention, the probability of committing a Type II error is denoted by β. In environmental assessment, a Type II error would be a “false negative” in which, for example, there is an effect of a pollutant on human health, but it is not detected. 20

We have followed standard statistical treatments that equate the calculated P-value with the estimate of Type I error rate α. However, Fisher’s evidential P-value may not be strictly equivalent to Neyman and Pearson’s α. Statisticians disagree whether the distinction is an important philosophical issue or simply a semantic difference. Hubbard and Bayarri (2003) argue that the incompatibility is important, and their paper is followed by discussion, comments, and rebuttals from other statisticians. Stay tuned! 20

The relationship between Type I and Type II errors informs discussions of the precautionary principle of environmental decision making. Historically, for example, regulatory agencies have assumed that new chemical products were benign until proven harmful. Very strong evidence was required to reject the null hypothesis of no effect on health and well-being. Manufacturers of chemicals and other potential pollutants are keen to minimize the probability of committing a Type I error. In contrast, environmental groups that serve the general public are interested in minimizing the probability that the manufacturer committed a Type II error. Such groups assume that a chemical is harmful until proven benign, and are willing to accept a larger probability of committing a Type I error if this means they can be more confident that the manufacturer has not falsely accepted the null hypothesis. Following such reasoning, in assessing quality control of industrial production, Type I and Type II errors are often known as producer and consumer errors, respectively (Sokal and Rohlf 1995).

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A concept related to the probability of committing a Type II error is the power of a statistical test. Power is calculated as 1 – β, and equals the probability of correctly rejecting the null hypothesis when it is false. We want our statistical tests to have good power so that we have a good chance of detecting significant patterns in our data when they are present. Ideally, we would like to minimize both Type I and Type II errors in our statistical inference. However, strategies designed to reduce Type I error inevitably increase the risk of Type II error, and vice versa. For example, suppose you decide to reject the null hypothesis only if P < 0.01—a fivefold more stringent standard than the conventional criterion of P < 0.05. Although your risk of committing a Type I error is now much lower, there is a much greater chance that when you fail to reject the null hypothesis, you may be doing so incorrectly (i.e., you will be committing a Type II error). Although Type I and Type II errors are inversely related to one another, there is no simple mathematical relationship between them, because the probability of a Type II error depends in part on what the alternative hypothesis is, how large an effect we hope to detect (Figure 4.5), the sample size, and the wisdom of our experimental design or sampling protocol.

WHAT IS THE RELATIONSHIP BETWEEN TYPE I AND TYPE II ERROR?

WHY ARE STATISTICAL DECISIONS BASED ON TYPE I ERROR? In contrast to the probability of committing a Type I error, which we determine with standard statistical tests, the probability of committing a Type II error is not often calculated or reported, and in many scientific papers, the probability of committing a Type II error is not even discussed. Why not? To begin with, we often cannot calculate the probability of a Type II error unless the alternative hypotheses are completely specified. In other words, if we want to determine the risk of falsely accepting the null hypothesis, the alternatives have to be fleshed out more than just “not H0.” In contrast, calculating the probability of a Type I error does not require this specification; instead we are required only to meet some assumptions of normality and independence (see Chapters 9 and 10). On a philosophical basis, some authors have argued that a Type I error is a more serious mistake in science than a Type II error (Shrader-Frechette and McCoy 1992). A Type I error is an error of falsity, in which we have incorrectly rejected a null hypothesis and made a claim about a more complex mechanism. Others may follow our work and try to build their own studies based on that false claim. In contrast, a Type II error is an error of ignorance. Although we have not rejected the null hypothesis, someone else with a better experiment or more data may be able to do so in the future, and the science will progress

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(A)

Statistical power (1 – β)

1.0 n = 50 n = 40 n = 30

0.8 0.6

n = 20 0.4 n = 10 n=5

0.2 0.0 0.005

0.01

0.025 P-value

0.05

0.1

(B) n = 50 n = 40 n = 30 n = 20

Statistical power (1 – β)

1.0 0.8

n = 10 0.6 0.4

Figure 4.5 The relationship between statistical power, P-values, and observable effect sizes as a function of sample size. (A) The P-value is the probability of incorrectly rejecting a true null hypothesis, whereas statistical power is the probability of correctly rejecting a false null hypothesis. The general result is that the lower the P-value used for rejection of the null hypothesis, the lower the statistical power of correctly detecting a treatment effect. At a given P-value, statistical power is greater when the sample size is larger. (B) The smaller the observable effect of the treatment (i.e., the smaller the difference between the treatment group and the control group), the larger the sample size necessary for good statistical power to detect a treatment effect.21

n=5

0.2 0.0 –100

–50 0 50 Difference between population means

100

from that point. However, in many applied problems, such as environmental monitoring or disease diagnosis, Type II errors may have more serious consequences because diseases or adverse environmental effects would not be correctly detected. 21

We can apply these graphs to our example comparing glucocorticoid hormone levels for populations of wolves exposed to snowmobiles (treatment group) versus wolves that were not exposed to snowmobiles (control group). In the original data (Creel et al. 2002), the standard deviation of the control population of wolves unexposed to snowmobiles was 73.1, and that of wolves exposed to snowmobiles was 114.2. Panel (A) suggests that if there were a 50-ng/g difference between the experimental populations, and if sample size was n = 50 in each group, the experimenters would have correctly accepted the alternative hypothesis only 51% of the time for P = 0.01. Panel (B) shows that power increases steeply as the populations become more different. In the actual well-designed study (Creel et al. 2002), sample size was 193 in the unexposed group and 178 in the exposed group, the difference between population means was 598 ng/g, and the actual power of the statistical test was close to 1.0 for P = 0.01.

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Hypothesis testing

Hypothesis A

Hypothesis B

Parameter estimation

B D

Hypothesis C

Hypothesis D

Figure 4.6 Hypothesis testing versus parameter estimation. Parameter estimation more easily accommodates multiple mechanisms and may allow for an estimate of the relative importance of different factors. Parameter estimation may involve the construction of confidence or credibility intervals (see Chapter 3) to estimate the strength of an effect. A related technique in the analysis of variance is to decompose the total variation in the data into proportions that are explained by different factors in the model (see Chapter 10). Both methods quantify the relative importance of different factors, whereas hypothesis testing emphasizes a binary yes/no decision as to whether a factor has a measurable effect or not.

Parameter Estimation and Prediction All the methods for hypothesis testing that we have described—the inductive method (and its modern descendant, Bayesian inference), the hypotheticodeductive method, and statistical hypothesis testing are concerned with choosing a single explanatory “answer” from an initial set of multiple hypotheses. In ecology and environmental science, it is more likely that many mechanisms may be operating simultaneously to produce observed patterns; an hypothesis-testing framework that emphasizes single explanations may not be appropriate. Rather than try to test multiple hypotheses, it may be more worthwhile to estimate the relative contributions of each to a particular pattern. This approach is sketched in Figure 4.6, in which we partition the effects of each hypothesis on the observed patterns by estimating how much each cause contributes to the observed effect. In such cases, rather than ask whether a particular cause has some effect versus no effect (i.e., is it significantly different from 0.0?), we ask what is the best estimate of the parameter that expresses the magnitude of the effect.22 For example, in Figure 4.3, measured photosynthetic rates for young sun leaves of 22 Chapters 9, 13, and 14 will introduce some of the strategies used for fitting curves and estimating parameters from data. See Hilborn and Mangel (1997) for a detailed discussion. Clark et al. (2003) describe Bayesian strategies to curve fitting.

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Summary

Rhizophora mangle were fit to a Michaelis-Menten equation. This equation is a simple model that describes a variable rising smoothly to an asymptote. The Michaelis-Menten equation shows up frequently in biology, being used to describe everything from enzyme kinematics (Real 1977) to invertebrate foraging rates (Holling 1959). The Michaelis-Menten equation takes the form Y=

kX X +D

where k and D are the two fitted parameters of the model, and X and Y are the independent and dependent variables. In this example, k represents the asymptote of the curve, which in this case is the maximum assimilation rate; the independent variable X is light intensity; and the dependent variable Y is the net assimilation rate. For the data in Figure 4.3, the parameter estimate for k is a maximum assimilation rate of 7.1 μmol CO2 m–2 s–1. This accords well with an “eyeball estimate” of where the asymptote would be on this graph. The second parameter in the Michaelis-Menten equation is D, the halfsaturation constant. This parameter gives the value of the X variable that yields a Y variable that is half of the asymptote. The smaller D is, the more quickly the curve rises to the asymptote. For the data in Figure 4.3, the parameter estimate for D is photosynthetically active radiation (PAR) of 250 μmol CO2 m–2 s–1. We also can measure the uncertainty in these parameter estimates by using estimates of standard error to construct confidence or credibility intervals (see Chapter 3). The estimated standard error for k = 0.49, and for D = 71.3. Statistical hypothesis testing and parameter estimation are related, because if the confidence interval of uncertainty includes 0.0, we usually are not able to reject the null hypothesis of no effect for one of the mechanisms. For the parameters k and D in Figure 4.3, the P-values for the test of the null hypothesis that the parameter does not differ from 0.0 are 0.0001 and 0.004, respectively. Thus, we can be fairly confident in our statement that these parameters are greater than 0.0. But, for the purposes of evaluating and fitting models, the numerical values of the parameters are more informative than just asking whether they differ or not from 0.0. In later chapters, we will give other examples of studies in which model parameters are estimated from data.

Summary Science is done using inductive and hypothetico-deductive methods. In both methods, observed data are compared with data predicted by the hypotheses. Through the inductive method, which includes modern Bayesian analyses, a

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single hypothesis is repeatedly tested and modified; the goal is to confirm or assert the probability of a particular hypothesis. In contrast, the hypothetico-deductive method requires the simultaneous statement of multiple hypotheses. These are tested against observations with the goal of falsifying or eliminating all but one of the alternative hypotheses. Statistics are used to test hypotheses objectively, and can be used in both inductive and hypothetico-deductive approaches. Probabilities are calculated and reported with virtually all statistical tests. The probability values associated with statistical tests may allow us to infer causes of the phenomena that we are studying. Tests of statistical hypotheses using the hypothetico-deductive method yield estimates of the chance of obtaining a result equal to or more extreme than the one observed, given that the null hypothesis is true. This P-value is also the probability of incorrectly rejecting a true null hypothesis (or committing a Type I statistical error). By convention and tradition, 0.05 is the cutoff value in the sciences for claiming that a result is statistically significant. The calculated P-value depends on the number of observations, the difference between the means of the groups being compared, and the amount of variation among individuals within each group. Type II statistical errors occur when a false null hypothesis is incorrectly accepted. This kind of error may be just as serious as a Type I error, but the probability of Type II errors is reported rarely in scientific publications. Tests of statistical hypotheses using inductive or Bayesian methods yield estimates of the probability of the hypothesis or hypotheses of interest given the observed data. Because these are confirmatory methods, they do not give probabilities of Type I or Type II errors. Rather, the results are expressed as the odds or likelihood that a particular hypothesis is correct. Regardless of the method used, all science proceeds by articulating testable hypotheses, collecting data that can be used to test the predictions of the hypotheses, and relating the results to underlying cause-and-effect relationships.

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CHAPTER 5

Three Frameworks for Statistical Analysis

In this chapter, we introduce three major frameworks for statistical analysis: Monte Carlo analysis, parametric analysis, and Bayesian analysis. In a nutshell, Monte Carlo analysis makes minimal assumptions about the underlying distribution of the data. It uses randomizations of observed data as a basis for inference. Parametric analysis assumes the data were sampled from an underlying distribution of known form, such as those described in Chapter 2, and estimates the parameters of the distribution from the data. Parametric analysis estimates probabilities from observed frequencies of events and uses these probabilities as a basis for inference. Hence, it is a type of frequentist inference. Bayesian analysis also assumes the data were sampled from an underlying distribution of known form. It estimates parameters not only from the data, but also from prior knowledge, and assigns probabilities to these parameters. These probabilities are the basis for Bayesian inference. Most standard statistics texts teach students parametric analysis, but the other two are equally important, and Monte Carlo analysis is actually easier to understand initially. To introduce these methods, we will use each of them to analyze the same sample problem.

Sample Problem Imagine you are trying to compare the nest density of ground-foraging ants in two habitats—field and forest. In this sample problem, we won’t concern ourselves with the scientific hypothesis that you are testing (perhaps you don’t even have one at this point); we will simply follow through the process of gathering and analyzing the data to determine whether there are consistent differences in the density of ant nests in the two habitats. You visit a forest and an adjacent field and estimate the average density of ant nests in each, using replicated sampling. In each habitat, you choose a random location, place a square quadrat of 1-m2 area, and carefully count all of the ant nests that occur within the quadrat. You repeat this procedure several times in

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Each row is an independent observation. The first column identifies the replicate with a unique ID number, the second column indicates the habitat sampled, and the third column gives the number of ant nests recorded in the replicate.

TABLE 5.1 Sample dataset used to illustrate Monte Carlo, parametric, and Bayesian analyses ID number

Habitat

Number of ant nests per quadrat

1 2 3 4 5 6 7 8 9 10

Forest Forest Forest Forest Forest Forest Field Field Field Field

9 6 4 6 7 10 12 9 12 10

each habitat. The issue of choosing random locations is very important for any type of statistical analysis. Without more complicated methods, such as stratified sampling, randomization is the only safeguard to ensure that we have a representative sample from a population (see Chapter 6). The spatial scale of the sampling determines the scope of inference. Strictly speaking, this sampling design will allow you to discuss differences between forest and field ant densities only at this particular site. A better design would be to visit several different forests and fields and sample one quadrat within each of them. Then the conclusions could be more readily generalized. Table 5.1 illustrates the data in a spreadsheet. The data are arranged in a table, with labeled rows and columns. Each row of the table contains all of the information on a particular observation. Each column of the table indicates a different variable that has been measured or recorded for each observation. In this case, your original intent was to sample 6 field and 6 forest quadrats, but the field quadrats were more time-consuming than you expected and you only managed to collect 4 field samples. Thus the data table has 10 rows (in addition to the first row, which displays labels) because you collected 10 different samples (6 from the forest and 4 from the field). The table has 3 columns. The first column contains the unique ID number assigned to each replicate. The other columns contain the two pieces of information recorded for each replicate: the habitat in which the replicate was sampled (field or forest); and the number of ant nests recorded in the quadrat.1 1

Many statistics texts would show these data as two columns of numbers, one for forest and one for field. However, the layout we have shown here is the one that is recognized most commonly by statistical software for data analysis.

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TABLE 5.2 Summary statistics for the sample data in Table 5.1 Habitat

Mean

Standard deviation

Forest Field

6 4

7.00 10.75

2.19 1.50

Following the procedures in Chapter 3, calculate the mean and standard deviation for each sample (Table 5.2). Plot the data, using a conventional bar chart of means and standard deviations (Figure 5.1), or the more informative box plot described in Chapter 3 (Figure 5.2). Although the numbers collected in the forest and field show some overlap, they appear to form two distinct groups: the forest samples with a mean of 7.0 nests per quadrat, and the field samples with a mean of 10.75 nests per quadrat. On the other hand, our sample size is very small (n = 6 forest and n = 4 field samples). Perhaps these differences could have arisen by chance or random sampling. We need to conduct a statistical test before deciding whether these differences are significant or not.

Monte Carlo Analysis Monte Carlo analysis involves a number of methods in which data are randomized or reshuffled so that observations are randomly reassigned to differ-

Ant nests/sample

Figure 5.1 Standard bar chart for sample data in

Field

Forest Habitat

Table 5.1. The height of the bar is the mean of the sample and the vertical line indicates one standard deviation above the mean. Monte Carlo, parametric, and Bayesian analyses can all be used to evaluate the difference in means between the groups.

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CHAPTER 5 Three Frameworks for Statistical Analysis

Figure 5.2 Box plot of data in Table 5.1. In a box plot, the central line within the box is the median of the data. The box includes 50% of the data. The top of the box indicates the 75th percentile (upper quartile) of the data, and the bottom of the box indicates the 25th percentile (lower quartile) of the data. The vertical lines extend to the upper and lower deciles (90th and 10th percentiles). For the field sample, there are so few data that the 75th and 90th percentiles do not differ. When data have asymmetric distributions or outliers, box plots may be more informative than standard bar graphs such as Figure 5.1.

Ant nests/sample

110

0 Field

Forest Habitat

ent treatments or groups. This randomization2 specifies the null hypothesis under consideration: that the pattern in the data is no different from that which we would expect if the observations were assigned randomly to the different groups. There are four steps in Monte Carlo analysis: 1. Specify a test statistic or index to describe the pattern in the data. 2. Create a distribution of the test statistic that would be expected under the null hypothesis. 3. Decide on a one- or two-tailed test. 4. Compare the observed test statistic to a distribution of simulated values and estimate the appropriate P-value as a tail probability (as described in Chapter 3). 2 Some statisticians distinguish Monte Carlo methods, in which samples are drawn from a known or specified statistical distribution, from randomization tests, in which existing data are reshuffled but no assumptions are made about the underlying distribution. In this book, we use Monte Carlo methods to mean randomization tests. Another set of methods includes bootstrapping, in which statistics are estimated by repeatedly subsampling with replacement from a dataset. Still another set of methods includes jackknifing, in which the variability in the dataset is estimated by systematically deleting each observation and then recalculating the statistics (see Figure 9.8 and the section “Discriminant Analysis” in Chapter 12). Monte Carlo, of course, is the famous gambling resort city on the Riviera, whose citizens do not pay taxes and are forbidden from entering the gaming rooms.

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Monte Carlo Analysis

Step 1: Specifying the Test Statistic

For this analysis, we will use as a measure of pattern the absolute difference in the means of the forest and field samples, or DIF: DIFobs = |10.75 – 7.00| = 3.75 The subscript “obs” indicates that this DIF value is calculated for the observed data. The null hypothesis is that a DIFobs equal to 3.75 is about what would be expected by random sampling. The alternative hypothesis would be that a DIFobs equal to 3.75 is larger than would be expected by chance. Step 2: Creating the Null Distribution

Next, estimate what DIF would be if the null hypothesis were true. To do this, use the computer (or a deck of playing cards) to randomly reassign the forest and field labels to the dataset. In the randomized dataset, there will still be 4 field and 6 forest samples, but those labels (Field and Forest) will be randomly reassigned (Table 5.3). Notice that in the randomly reshuffled dataset, many of the observations were placed in the same group as the original data. This will happen by chance fairly often in small datasets. Next, calculate the sample statistics for this randomized dataset (Table 5.4). For this dataset, DIFsim = |7.75 – 9.00| = 1.25. The subscript “sim” indicates that this value of DIF is calculated for the randomized, or simulated, data. In this first simulated dataset, the difference between the means of the two groups (DIFsim = 1.25) is smaller than the difference observed in the real data (DIFobs = 3.75). This result suggests that means of the forest and field samples may differ more than expected under the null hypothesis of random assignment. TABLE 5.3 Monte Carlo randomization of the habitat labels in Table 5.1 Habitat

Number of ant nests per quadrat

Field Field Forest Forest Field Forest Forest Field Forest Forest

9 6 4 6 7 10 12 9 12 10

In the Monte Carlo analysis, the sample labels in Table 5.1 are reshuffled randomly among the different samples. After reshuffling, the difference in the means of the two groups, DIF, is recorded. This procedure is repeated many times, generating a distribution of DIF values (see Figure 5.3).

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TABLE 5.4 Summary statistics for the randomized data in Table 5.3 Habitat

Mean

Standard deviation

Forest Field

6 4

9.00 7.75

3.286 1.500

In the Monte Carlo analysis, these values represent the mean and standard deviation in each group after a single reshuffling of the labels. The difference between the two means (DIF = |7.75 – 9.00| = 1.25) is the test statistic.

However, there are many different random combinations that can be produced by reshuffling the sample labels.3 Some of these have relatively large values of DIFsim and some have relatively small values of DIFsim. Repeat this reshuffling exercise many times (usually 1000 or more), then illustrate the distribution of the simulated DIF values with a histogram (Figure 5.3) and summary statistics (Table 5.5). The mean DIF of the simulated datasets was only 1.46, compared to the observed value of 3.75 for the original data. The standard deviation of 2.07 could be used to construct a confidence interval (see Chapter 3), but it should not be, because the distribution of simulated values (see Figure 5.3) has a long righthand tail and does not follow a normal distribution. An approximate 95% confidence interval can be derived from this Monte Carlo analysis by identifying the upper and lower 2.5 percentiles of the data from the set of simulated DIF values and using these values as the upper and lower bounds of the confidence interval (Efron 1982). Step 3: Deciding on a One- or Two-Tailed Test

Next, decide whether to use a one-tailed test or a two-tailed test. The “tail” refers to the extreme left- or right-hand areas under the probability density function (see Figure 3.7). It is these tails of the distribution that are used to determine the cutoff points for a statistical test at P = 0.05 (or any other probability level). A one-tailed test uses only one tail of the distribution to estimate the P-value. A two-tailed test uses both tails of the distribution to estimate the P-value. In a

With 10 samples split into a group of 6 and a group of 4, there are

⎛10⎞ ⎜ ⎟ = 210 ⎝ 4⎠ combinations that can be created by reshuffling the labels (see Chapter 2). However, the number of unique values of DIF that are possible is somewhat less than this because some of the samples had identical nest counts.

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Monte Carlo Analysis

300

Number of simulations

250

200

150

Observed value(3.75)

100

0 1

DIF

Figure 5.3 Monte Carlo analysis of the data in Table 5.1. For each randomization, the data labels (Forest, Field) were randomly reshuffled among the replicates. Next, the difference (DIF) between the means of the two groups was calculated. This histogram illustrates the distribution of DIF values from 1000 such randomizations. The arrow indicates the single value of DIF observed in the real dataset (3.75). The observed DIF sits well in the right-hand tail of the distribution. The observed DIF of 3.75 was larger than or equal to all but 36 of the simulated DIF values. Therefore, under the null hypothesis of random assignment of samples to groups, the tail probability of finding this observation (or one more extreme) is 36/1000 = 0.036.

one-tailed test, all 5% of the area under the curve is located in one tail of the distribution. In a two-tailed test, each tail of the distribution would encompass 2.5% of the area. Thus, a two-tailed test requires more extreme values to achieve statistical significance than does a one-tailed test. However, the cutoff value is not the most important issue in deciding whether to use a one- or a two-tailed test. Most important are the nature of the response TABLE 5.5 Summary statistics for 1000 simulated values of DIF in Figure 5.3 Variable

DIFsim

Mean

Standard deviation

1000

1.46

2.07

In the Monte Carlo analysis, each of these values was created by reshuffling the data labels in Table 5.1, and calculating the difference between the means of the two groups (DIF). Calculation of the P-value does not require that DIF follow a normal distribution, because the P-value is determined directly by the location of the observed DIF statistic in the histogram (see Table 5.6).

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variable and the precise hypothesis being tested. In this case, the response variable was DIF, the absolute difference in the means of forest and field samples. For the DIF variable, a one-tailed test is most appropriate for unusually large values of DIF. Why not use a two-tailed test with DIF? The lower tail of the DIFsim distribution would represent values of DIF that are unusually small compared to the null hypothesis. In other words, a two-tailed test would also be testing for the possibility that forest and field means were more similar than expected by chance. This test for extreme similarity is not biologically informative, so attention is restricted to the upper tail of the distribution. The upper tail represents cases in which DIF is unusually large compared to the null distribution. How could the analysis be modified to use a two-tailed test? Instead of using the absolute value of DIF, use the average difference between forest and field samples (DIF*). Unlike DIF, DIF* can take on both positive and negative values. DIF* will be positive if the field average is greater than the forest average. DIF* will be negative if the field average is less than the forest average. As before, randomize the data and create a distribution of DIF*sim. In this case, however, a two-tailed test would detect cases in which the field mean was unusually large compared to the forest mean (DIF* positive) and cases in which the field mean was unusually small compared to the forest mean (DIF* negative). Whether you are using Monte Carlo, parametric, or Bayesian analysis, you should study carefully the response variable you are using. How would you interpret an extremely large or an extremely small value of the response variable relative to the null hypothesis? The answer to this question will help you decide whether a one- or a two-tailed test is most appropriate. Step 4: Calculating the Tail Probability

The final step is to estimate the probability of obtaining DIFobs or a value more extreme, given that the null hypothesis is true [P(data|H0)]. To do this, examine the set of simulated DIF values (plotted as the histogram in Figure 5.3), and tally up the number of times that the DIFobs was greater than, equal to, or less than each of the 1000 values of DIFsim. In 29 of 1000 randomizations, DIFsim = DIFobs, so the probability of obtaining DIFobs under the null hypothesis is 29/1000 = 0.029 (Table 5.6). However, when we calculate a statistical test, we usually are not interested in this exact probability as much as we are the tail probability. That is, we want to know the chances of obtaining an observation as large or larger than the real data, given that the null hypothesis is true. In 7 of 1000 randomizations, DIFsim > DIFobs. Thus, the probability that DIFobs ≥ 3.75 is (7 + 29)/1000 = 0.036. This tail probability is the frequency of obtaining the observed value (29/1000) plus the frequency of obtaining a more extreme result (7/1000).

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Monte Carlo Analysis

TABLE 5.6 Calculation of tail probabilities in Monte Carlo analysis Inequality

DIFsim > DIFobs DIFsim = DIFobs DIFsim < DIFobs

7 29 964

115

Comparisons of DIFobs (absolute difference in the mean of the two groups in the original data) with DIFsim (absolute difference in the mean of the two groups after randomizing the group assignments). N is the number of simulations (out of 1000) for which the inequality was obtained. Because DIFsim DIFobs in 7 + 29 = 36 trials out of 1000, the tail probability under the null hypothesis of finding DIFobs this extreme is 36/1000 = 0.036.

Follow the procedures and interpretations of P-values that we discussed in Chapter 4. With a tail probability of 0.036, it is unlikely that these data would have occurred given the null hypothesis is true. Assumptions of Monte Carlo Methods

Monte Carlo methods rest on three assumptions: 1. The data collected represent random, independent samples. 2. The test statistic describes the pattern of interest. 3. The randomization creates an appropriate null distribution for the question. Assumptions 1 and 2 are common to all statistical analyses. Assumption 1 is the most critical, but it is also the most difficult to confirm, as we will discuss in Chapter 6. Assumption 3 is easy to meet in this case. The sampling structure and null hypothesis being tested are very simple. For more complex questions, however, the appropriate randomization method may not be obvious, and there may be more than one way to construct the null distribution (Gotelli and Graves 1996). Advantages and Disadvantages of Monte Carlo Methods

The chief conceptual advantage of Monte Carlo methods is that it makes clear and explicit the underlying assumptions and the structure of the null hypothesis. In contrast, conventional parametric analyses often gloss over these features, perhaps because the methods are so familiar. Another advantage of Monte Carlo methods over parametric analysis is that it does not require the assumption that the data are sampled from a specified probability distribution, such as the normal. Finally, Monte Carlo simulations allow you to tailor your statistical test to particular questions and datasets, rather than having to shoehorn them into a conventional test that may not be the most powerful method for your question, or whose assumptions may not match the sampling design of your data. The chief disadvantage of Monte Carlo methods is that it is computer-intensive and is not included in most traditional statistical packages (but see Gotelli

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and Entsminger 2003). As computers get faster and faster, older limitations on computational methods are disappearing, and there is no reason that even very complex statistical analyses cannot be run as a Monte Carlo simulation. However, until such routines become widely available, Monte Carlo methods are available only to those who know a programming language and can write their own programs.4 A second disadvantage of Monte Carlo analysis is psychological. Some scientists are uneasy about Monte Carlo methods because different analyses of the same dataset can yield slightly different answers. For example, we re-ran the analysis on the ant data in Table 5.1 ten times and got P-values that ranged from 0.030 to 0.046. Most researchers are more comfortable with parametric analyses, which have more of an air of objectivity because the same P-value results each time the analysis is repeated. A final weakness is that the domain of inference for a Monte Carlo analysis is subtly more restrictive than that for a parametric analysis. A parametric analysis assumes a specified distribution and allows for inferences about the underlying parent population from which the data were sampled. Strictly speaking, inferences from Monte Carlo analyses (at least those based on simple randomization tests) are limited to the specific data that have been collected. However, if a sample is representative of the parent population, the results can be generalized cautiously.

4 One of the most important things you can do is to take the time to learn a real programming language. Although some individuals are proficient at programming macros in spreadsheets, macros are practical only for the most elementary calculations; for anything more complex, it is actually simpler to write a few lines of computer code than to deal with the convoluted (and error-prone) steps necessary to write macros. There are now many computer languages available for you to choose from. We both prefer to use R, an open-source package that includes many built-in mathematical and statistical functions (R-project.org). Unfortunately, learning to program is like learning to speak a foreign language—it takes time and practice, and there is no immediate payoff. Sadly, our academic culture doesn’t encourage the learning of programming skills (or languages other than English). But if you can overcome the steep learning curve, the scientific payoff is tremendous. The best way to begin is not to take a class, but to obtain software, work through examples in the manual, and try to code a problem that interests you. Hilborn and Mangel’s The Ecological Detective (1997) contains an excellent series of ecological exercises to build your programming skills in any language. Bolker (2008) provides a wealth of worked examples using R for likelihood analyses and stochastic simulation models. Not only will programming free you from the chains of canned software packages, it will sharpen your analytical skills and give you new insights into ecological and statistical models. You will have a deeper understanding of a model or a statistic once you have successfully programmed it!

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Parametric Analysis

Parametric Analysis Parametric analysis refers to the large body of statistical tests and theory built on the assumption that the data being analyzed were sampled from a specified distribution. Most statistical tests familiar to ecologists and environmental scientists specify the normal distribution. The parameters of the distribution (e.g., the population mean μ and variance σ2) are then estimated and used to calculate tail probabilities for a true null hypothesis. A large statistical framework has been built around the simplifying assumption of normality of data. As much as 80% to 90% of what is taught in standard statistics texts falls under this umbrella. Here we use a common parametric method, the analysis of variance (ANOVA), to test for differences in the group means of the sample data.5 There are three steps in parametric analysis: 1. Specify the test statistic. 2. Specify the null distribution. 3. Calculate the tail probability. Step 1: Specifying the Test Statistic

Parametric analysis of variance assumes that the data are drawn from a normal, or Gaussian, distribution. The mean and variance of these curves can be estimated from the sample data (see Table 5.1) using Equations 3.1 and 3.9. Figure 5.4 shows the distributions used in parametric analysis of variance. The original data are arrayed on the x-axis, and each color represents a different habitat (black circles for the forest samples, blue circles for the field samples).

The framework for modern parametric statistical theory was largely developed by the remarkable Sir Ronald Fisher (1890–1962), and the F-ratio is named in his honor (although Fisher himself felt that the ratio needed further study and refinement). Fisher held the Balfour Chair in Genetics at Cambridge University from 1943 to 1957 and made fundamental contributions to the theory of population genetics and evolution. In statistics, he developed the analysis of variance (ANOVA) to analyze crop yields in agricultural systems, in Sir Ronald Fisher which it may be difficult or impossible to replicate treatments. Many of the same constraints face ecologists in the design of their experiments today, which is why Fisher’s methods continue to be so useful. His classic book, The Design of Experiments (1935) is still enjoyable and worthwhile reading. It is ironic that Fisher became uneasy about his own methods when dealing with experiments he could design well, whereas today many ecologists apply his methods to ad hoc observations in poorly designed natural experiments (see Chapters 4 and 6). (Photograph courtesy of the Ronald Fisher Memorial Trust.)

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200 Null hypothesis 150 Frequency

118

Variation among groups 100 Variation within groups 50

0 0

15 10 Ant nest entrances/sample

Figure 5.4 Normal distributions based on sample data in Table 5.1 The data in Table 5.1 are shown as symbols (black circles and curve, forest samples; blue circles and curve, field samples) that indicate the number of ant nests counted in each quadrat. The null hypothesis is that all the data were drawn from the same population whose normal distribution is indicated by the dashed line. The alternative hypothesis is that each habitat has its own distinct mean (and variance), indicated by the two smaller normal distributions. The smaller the shaded overlap of the two distributions, the less likely it is that the null hypothesis is true. Measures of variation among and within groups are used to calculate the F-ratio and test the null hypothesis.

First consider the null hypothesis: that both sets of data were drawn from a single underlying normal distribution, which is estimated from the mean and variance of all the data (dashed curve; mean = 8.5, standard deviation = 2.54). The alternative hypothesis is that the samples were drawn from two different populations, each of which can be characterized by a different normal distributions, one for the forest and one for the field. Each distribution has its own mean, although we assume the variance is the same (or similar) for each of the two groups. These two curves are also illustrated in Figure 5.4, using the summary statistics calculated in Table 5.2. How is the null hypothesis tested? The closer the two curves are for the forest and field data, the more likely the data would be collected given the null hypothesis is true, and the single dashed curve best represents the data. Conversely, the more separate the two curves are, the less likely it is that the data represent a single population with a common mean and variance. The area of overlap between these two distributions (shaded in Figure 5.4) should be a measure of how close or how far apart the distributions are.

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Parametric Analysis

Fisher’s contribution was to quantify that overlap as a ratio of two variables. The first is the amount of variation among the groups, which we can think of as the variance (or standard deviation) of the means of the two groups. The second is the amount of variation within each group, which we can think of as the variance of the observations around their respective means. Fisher’s F-ratio can be interpreted as a ratio of these two sources of variation: F = (variance among groups + variance within groups) / variance within groups

(5.1)

In Chapter 10, we will explain in detail how to calculate the numerator and denominator of the F-ratio. For now, we simply emphasize that the ratio measures the relative size of two sources of variation in the data: variation among groups and within groups. For these data, the F-ratio is calculated as 33.75/ 3.84 = 8.78. In an ANOVA, the F-ratio is the test statistic that describes (as a single number) the pattern of differences among the means of the different groups being compared. Step 2: Specifying the Null Distribution

The null hypothesis is that all the data were drawn from the same population, so that any differences between the means of the groups are no larger than would be expected by chance. If this null hypothesis is true, then the variation among groups will be small, and we expect to find an F-ratio of 1.0. The F-ratio will be correspondingly larger than 1.0 if the means of the groups are widely separated (large among-group variation) relative to the variation within groups.6 In this example, the observed F-ratio of 8.78 is almost 10 times larger than the expected value of 1.0, a result that seems unlikely if the null hypothesis were true. Step 3: Calculating the Tail Probability

The P-value is an estimate of the probability of obtaining an F-ratio ≥ 8.78, given that the null hypothesis is true. Figure 5.5 shows the theoretical distribution of the F-ratio and the observed F-ratio of 8.78, which lies in the extreme right hand tail of the distribution. What is its tail probability (or P-value)?

As in Monte Carlo analysis, there is the theoretical possibility of obtaining an F-ratio that is smaller than expected by chance. In such a case, the means of the groups are unusually similar, and differ less than expected if the null hypothesis were true. Unusually small F-ratios are rarely seen in the ecological literature, although Schluter (1990) used them as an index of species-for-species matching of body sizes and community convergence.

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0.0008

0.0006 Probability density

120

0.0004

0.0002 Critical value: F = 5.32

Observed value: F = 8.78

0 0

7 8 F-ratio

Figure 5.5 Theoretical F-distribution. The larger the observed F-ratio, the more unlikely it would be if the null hypothesis were true. The critical value for this distribution equals 5.32; the area under the curve beyond this point is equal to 5% of the area under the entire curve. The observed F-ratio of 8.78 lies beyond the critical value, so P(ant data | null hypothesis) ≤ 0.05. In fact, the actual probability of P(ant data | null hypothesis) = 0.018, because the area under the curve to the right of the observed F-ratio represents 1.8% of the total area under the curve. Compare this result to the P-value of 0.036 from the Monte Carlo analysis (see Figure 5.3).

The P-value of this F-ratio is calculated as the probability mass of the F-ratio distribution (or area under the curve) equal to or greater than the observed Fratio. For these data, the probability of obtaining an F-ratio as large or larger than 8.78 (given two groups and a total N of 10) equals 0.018. As the P-value is smaller than 0.05, we consider it unlikely that such a large F-ratio would occur by chance alone, and we reject the null hypothesis that our data were sampled from a single population. Assumptions of the Parametric Method

There are two assumptions for all parametric analyses: 1. The data collected represent random, independent samples. 2. The data were sampled from a specified distribution. As we noted for Monte Carlo analysis, the first assumption of random, independent sampling is always the most important in any analysis. The second assumption is usually satisfied because normal (bell-shaped) distributions are ubiquitous and turn up frequently in the real world.

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Specific parametric tests usually include additional assumptions. For example, ANOVA further assumes that variances within each group being compared are equal (see Chapter 10). If sample sizes are large, this assumption can be modestly violated and the results will still be robust. However, if sample sizes are small (as in this example), this assumption is more important. Advantages and Disadvantages of the Parametric Method

The advantage of the parametric method is that it uses a powerful framework based on known probability distributions. The analysis we presented was very simple, but there are many parametric tests appropriate for complex experimental and sampling designs (see Chapter 7). Although parametric analysis is intimately associated with the testing of statistical null hypotheses, it may not be as powerful as sophisticated Monte Carlo models that are tailored to particular questions or data. In contrast to Bayesian analysis, parametric analysis rarely incorporates a priori information or results from other experiments. Bayesian analysis will be our next topic— after a brief detour into non-parametric statistics. Non-Parametric Analysis: A Special Case of Monte Carlo Analysis

Non-parametric statistics are based on the analysis of ranked data. In the ant example, we would rank the observations from largest to smallest and then calculate statistics based on the sums, distributions, or other synthetic measures of those ranks. Non-parametric analyses do not assume a specified parametric distribution (hence the name), but they still require independent, random sampling, (as do all statistical analyses). A non-parametric test is in essence a Monte Carlo analysis of ranked data, and non-parametric statistical tables give P-values that would be obtained by a randomization test on ranks. Thus, we have already described the general rationale and procedures for such tests. Although they are used commonly by some ecologists and environmental scientists, we do not favor non-parametric analyses for three reasons. First, using ranked data wastes information that is present in the original observations. A Monte Carlo analysis of the raw data is much more informative, and often more powerful. One justification that is offered for a non-parametric analysis is that the ranked data may be more robust to measurement error. However, if the original observations are so error-prone that only the ranks are reliable, it is probably a good idea to re-do the experiment using measurement methods that offer greater accuracy. Second, relaxing the assumption of a parametric distribution (e.g., normality) is not such a great advantage, because parametric analyses often are robust to violations of this assumption (thanks to the Central Limit Theorem). Third, non-parametric methods are available only for extremely simple

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experimental designs, and cannot easily incorporate covariates or blocking structures. We have found that virtually all needs for statistical analysis of ecological and environmental data can be met by parametric, Monte Carlo, or Bayesian approaches.

Bayesian Analysis Bayesian analysis is the third major framework for data analysis. Scientists often believe that their methods are “objective” because they treat each experiment as a tabula rasa (blank slate): the simple statistical null hypothesis of random variation reflects ignorance about cause and effect. In our example of ant nest densities in forests and fields, our null hypothesis is that the two are equal, or that being in forests and fields has no consistent effect on ant nest density. Although it is possible that no one has ever investigated ant nests in forests and fields before, it is extremely unlikely; our reading of the literature on ant biology prompted us to conduct this particular study. So why not use data that already exist to frame our hypotheses? If our only goal is the hypothetico-deductive one of falsification of a null hypothesis, and if previous data all suggest that forests and fields differ in ant nest densities, it is very likely that we, too, will falsify the null hypothesis. Thus, we needn’t waste time or energy doing the study yet again. Bayesians argue that we could make more progress by specifying the observed difference (e.g., expressed as the DIF or the F-ratio described in the previous sections), and then using our data to extend earlier results of other investigators. Bayesian analysis allows us to do just this, as well as to quantify the probability of the observed difference. This is the most important difference between Bayesian and frequentist methods. There are six steps in Bayesian inference: 1. 2. 3. 4. 5. 6.

Specify the hypothesis. Specify parameters as random variables. Specify the prior probability distribution. Calculate the likelihood. Calculate the posterior probability distribution. Interpret the results.

Step 1: Specifying the Hypothesis

The primary goal of a Bayesian analysis is to determine the probability of the hypothesis given the data that have been collected: P(H | data). The hypothesis needs to be quite specific, and it needs to be quantitative. In our parametric analysis of ant nest density, the hypothesis of interest (i.e., the alternative hypothesis)

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was that the samples were drawn from two populations with different means and equal variances, one for the forest and one for the field. We did not test this hypothesis directly. Instead, we tested the null hypothesis: the observed value of F was no larger than that expected if the samples were drawn from a single population. We found that the observed F-ratio was improbably large (P = 0.018), and we rejected the null hypothesis. We could specify more precisely the null hypothesis and the alternative hypothesis as hypotheses about the value of the F-ratio. Before we can specify these hypotheses, we need to know the critical value for the F-distribution in Figure 5.5. In other words, how large does an F-ratio have to be in order to have a P-value ≤ 0.05? For 10 observations of ants in two groups (field and forest), the critical value of the F-distribution (i.e., that value for which the area under the curve equals 5% of the total area) equals 5.32 (see Figure 5.5). Thus, any observed F-ratio greater than or equal to 5.32 would be grounds for rejecting the null hypothesis. Remember that the general hypothetico-deductive statement of the probability of the null hypothesis is P(data | H0). In the ant nest example, the data result in an F-ratio equal to 8.78. If the null hypothesis is true, the observed Fratio should be a random sample from the F-distribution shown in Figure 5.5. Therefore we ask what is P(Fobs = 8.78 | Ftheoretical)? In contrast, Bayesian analysis proceeds by inverting this probability statement: what is the probability of the hypothesis given the data we collected [P(H | data)]? The ant nest data can be expressed as F = 8.78. How are the hypotheses expressed in terms of the F-distribution? The null hypothesis is that the ants were sampled from a single population. In this case, the expected value of the F-ratio is small (F < 5.32, the critical value). The alternative hypothesis is that the ants were sampled from two populations, in which case the F-ratio would be large (F = 5.32). Therefore, the Bayesian analysis of the alternative hypothesis calculates P(F 5.32 | Fobs = 8.78). By the First Axiom of Probability, P(F < 5.32 | Fobs) = 1 – P(F 5.32 | Fobs). A modification of Bayes’ Theorem (introduced in Chapter 1) allows us to directly calculate P(hypothesis | data): P(hypothesis | data) =

P(hypothesis)P(data | hypothesis) P(data)

(5.2)

In Equation 5.2, P(hypothesis | data) on the left-hand side of the equation is called the posterior probability distribution (or simply the posterior), and is the quantity of interest. The right-hand side of the equation consists of a fraction. In the numerator, the term P(hypothesis) is referred to as the prior

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probability distribution (or simply the prior), and is the probability of the hypothesis of interest before you conducted the experiment. The next term in the numerator, P(data | hypothesis), is referred to as the likelihood of the data; it reflects the probability of observing the data given the hypothesis.7 The denominator, P(data) is a normalizing constant that reflects the probability of the data given all possible hypotheses.8 Because it is simply a normalizing constant (and so scales our posterior probability to the range [0,1]), P(hypothesis | data) ∝ P(hypothesis)P(data | hypothesis) (where ∝ means “is proportional to”) and we can focus our attention on the numerator. Returning to the ants and their F-ratios, we focus on P(F 5.32 | Fobs = 8.78). We have now specified our hypothesis quantitatively in terms of the relationship between the F-ratio we observe (the data) and the critical value of F 5.32 (the hypothesis). This is a more precise hypothesis than the hypothesis dis-

7 Fisher developed the concept of likelihood as a response to his discomfort with Bayesian methods of inverse probability:

What has now appeared, is that the mathematical concept of probability is inadequate to express our mental confidence or diffidence in making such inferences, and that the mathematical quantity which appears to be appropriate for measuring our order of preference among different possible populations does not in fact obey the laws of probability. To distinguish it from probability, I have used the term ‘Likelihood’ to designate this quantity. (Fisher 1925, p. 10). The likelihood is written as L(hypothesis | data) and is directly proportional (but not equal to) the probability of the observed data given the hypothesis of interest: L(hypothesis | data) = cP(dataobs | hypothesis). In this way, the likelihood differs from a frequentist P-value, because the P-value expresses the probability of the infinitely many possible samples of the data given the statistical null hypothesis (Edwards 1992). Likelihood is used extensively in information-theoretic approaches to statistical inference (e.g., Hilborn and Mangel 1997; Burnham and Anderson 2010), and it is a central part of Bayesian inference. However, likelihood does not follow the axioms of probability. Because the language of probability is a more consistent way of expressing our confidence in a particular outcome, we feel that statements of the probabilities of different hypotheses (which are scaled between 0 and 1) are more easily interpreted than likelihoods (which are not). 8

The denominator is calculated as

∫ P(H i ) P(data | H i )dH i

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cussed in the previous two sections, that there is a difference between the density of ants in fields and in forests. Step 2: Specifying Parameters as Random Variables

A second fundamental difference between frequentist analysis and Bayesian analysis is that, in a frequentist analysis, parameters (such as the true population means μforest and μfield, their standard deviations σ2, or the F-ratio) are fixed. In other words, we assume there is a true value for the density of ant nests in fields and forests (or at least in the field and forest that we sampled), and we estimate those parameters from our data. In contrast, Bayesian analysis considers these parameters to be random variables, with their own associated parameters (e.g., means, variances). Thus, for example, instead of the population mean of ant colonies in the field being a fixed value μfield, the mean could be expressed as a normal random variable with its own mean and variance: μfield ∼ N(λfield,σ2). Note that the random variable representing ant colonies does not have to be normal. The type of random variable used for each population parameter should reflect biological reality, not statistical or mathematical convenience. In this example, however, it is reasonable to describe ant colony densities as normal random variables: μfield ∼ N(λfield,σ2), μforest ∼ N(λforest,σ2). Step 3: Specifying the Prior Probability Distribution

Because our parameters are random variables, they have associated probability distributions. Our unknown population means (the λfield and λforest terms) themselves have normal distributions, with associated unknown means and variances. To do the calculations required by Bayes’ Theorem, we have to specify the prior probability distributions for these parameters—that is, what are probability distributions for these random variables before we do the experiment?9

The specification of priors is a fundamental division between frequentists and Bayesians. To a frequentist, specifying a prior reflects subjectivity on the part of the investigator, and thus the use of a prior is considered unscientific. Bayesians argue that specifying a prior makes explicit all the hidden assumptions of an investigation, and so it is a more honest and objective approach to doing science. This argument has lasted for centuries (see reviews in Effron 1986 and in Berger and Berry 1988), and was one reason for the marginalization of Bayesians within the statistical community. However, the advent of modern computational techniques allowed Bayesians to work with uninformative priors, such as the ones we use here. It turns out that, with uninformative priors, Bayesian and frequentist results are very similar, although their final interpretations remain different. These findings have led to a recent renaissance in Bayesian statistics and relatively easy-to-use software for Bayesian calculations is now widely available (Kéry 2010).

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We have two basic choices for specifying the prior. First, we can comb and reanalyze data in the literature, talk to experts, and come up with reasonable estimates for the density of ant nests in fields and forests. Alternatively, we can use an uninformative prior, for which we initially estimate the density of ant nests to be equal to zero and the variances to be very large. (In this example, we set the population variances to be equal to 100,000.) Using an uninformative prior is equivalent to saying that we have no prior information, and that the mean could take on virtually any value with roughly equal probability.10 Of course, if you have more information, you can be more specific with your prior. Figure 5.6 illustrates the uninformative prior and a (hypothetical) more informative one. Similarly, the standard deviation σ for μfield and μforest also has a prior probability distribution. Bayesian inference usually specifies an inverse gamma distribution11 for the variances; as with the priors on the means, we use an uninformative prior for the variance. We write this symbolically as σ2 ∼ IG(1,000, 1,000) (read “the variance is distributed as an inverse gamma distribution with parameters 1,000 and 1,000). We also calculate the precision of our estimate of variance, which we symbolize as τ, where τ = 1/σ2. Here τ is a gamma random variable (the inverse of an inverse gamma is a gamma), and we write this symbolically as τ ∼ Γ(0.001, 0.001) (read “tau is distributed as a gamma distribution with parameters 0.001, 0.001”). The form of this distribution is illustrated in Figure 5.7.

You might ask why we don’t use a uniform distribution, in which all values have equal probability. The reason is that the uniform distribution is an improper prior. Because the integral of a uniform distribution is undefined, we cannot use it to calculate a posterior distribution using Bayes’ Theorem. The uninformative prior N(0,100,000) is nearly uniform over a huge range, but it can be integrated. See Carlin and Louis (2000) for further discussion of improper and uninformative priors. 11

The gamma distribution for precision and the inverse gamma distribution for variance are used for two reasons. First, the precision (or variance) needs to take on only positive values. Thus, any probability density function that has only positive values could be used for priors for precision or variance. For continuous variables, such distributions include a uniform distribution that is restricted to positive numbers and the gamma distribution. The gamma distribution is somewhat more flexible than the uniform distribution and allows for better incorporation of prior knowledge. Second, before the use of high-speed computation, most Bayesian analyses were done using conjugate analysis. In a conjugate analysis, a prior probability distribution is sought that has the same form as the posterior probability distribution. This convenient mathematical property allows for closed-form (analytical) solutions to the complex integration involved in Bayes’ Theorem. For data that are normally distributed, the conjugate prior for the parameter that specifies the mean is a normal distribution, and the conjugate prior for the parameter that specifies the precision (= 1/variance) is a gamma distribution. For data that are drawn from a Poisson distribution, the gamma distribu-

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0.004

N~(0, 100)

Probability

0.003

0.002

0.001 N~(0, 100,000) 0.000 –4000

–2000 0 Average λ ant density (λ)

2000

4000

Figure 5.6 Prior probability distributions for Bayesian analysis. Bayesian analysis requires specification of prior probability distributions for the statistical parameters of interest. In this analysis of the data in Table 5.1, the parameter is average ant density, λ. We begin with a simple uninformative prior probability distribution that average ant density is described by a normal distribution with mean 0 and standard deviation of 100,000 (blue curve). Because the standard deviation is so large, the distribution is nearly uniform over a large range of values: between –1500 and +1500, the probability is essentially constant (∼0.0002), which is appropriate for an uninformative prior. The black curve represents a more precise prior probability distribution. Because the standard deviation is much smaller (100), the probability is no longer constant over a larger range of values, but instead decreases more sharply at extreme values.

tion is the conjugate prior for the parameter that defines the mean (or rate) of the Poisson distribution. For further discussion, see Gelman et al. (1995). The gamma distribution is a two-parameter distribution, written as Γ(a,b), where a is referred to the shape parameter and b is the scale parameter. The probability density function of the gamma distribution is

P( X ) =

b a (a −1) – bX X e , for X > 0 Γ(a)

where Γ(a) is the gamma function Γ(n) = (n – 1)! for integers n > 0. More generally, for real numbers z, the gamma function is defined as z −1

⎡ 1⎤ ∫ 0 ⎢⎣ln t ⎥⎦ dt The gamma distribution has expected value E(X) = a/b and variance = a/b2. Two distributions used commonly by statisticians are special cases of the gamma distribution. The χ2 distribution with ν degrees of freedom is equal to Γ(ν/2, 0.5). The exponential distribution with parameter β that was discussed in Chapter 2 is equal to Γ(1,β). Finally, if the random variable 1/X ∼ Γ(a,b), then X is said to have an inverse gamma (IG) distribution. To obtain an uninformative prior for the variance of a normal random variable, we take the limit of the IG distribution as a and b both approach 0. This is the reason we use a = b = 0.001 as the prior parameters for the gamma distribution describing the precision of the estimate. Γ(z ) =

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This use of precision should make some intuitive sense; because our estimate of variability decreases when we have more information, the precision of our estimate increases. Thus, a high value of precision equals a low variance of our estimated parameter. We now have our prior probability distributions. The unknown population means of ants in fields and forests, μfield and μforest, are both normal random variables with expected values equal to λfield and λforest and unknown (but equal) variances, σ2. These expected means themselves are normal random variables with expected values of 0 and variances of 100,000. The population precision τ is the reciprocal of the population variance σ2, and is a gamma random variable with parameters (0.001, 0.001): μi ~ N(λi, σ2) λi ~ N(0, 100,000) τ = 1/σ2 ~ Γ(0.001, 0.001) These equations completely specify P(hypothesis) in the numerator of Equation 5.2. If we had real prior information, such as the density of ant nests in other fields and forests, we could use those values to more accurately specify the expected means and variances of the λ’s.

0.004

Figure 5.7 Uninformative prior

0.003

Probability

probability distribution for the precision (= 1/variance). Bayesian inference requires not only a specification of a prior distribution for the mean of the variable (see Figure 5.6), but also a specification for the precision (= 1/variance). Bayesian inference usually specifies an inverse gamma distribution for the variances. As with the distribution of the means, an uninformative prior is used for the variance. In this case, the variance is distributed as an inverse gamma distribution with parameters 1000 and 1000: σ2 ∼ IG(1,000, 1,000). Because the variance is very large, the precision is small.

0.002

0.001

0 0.05

0.30

0.55 0.80 1.05 Precision (= 1/σ2)

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Step 4: Calculating the Likelihood

The other quantity in the numerator of Bayes’ Theorem (see Equation 5.2) is the likelihood, P(dataobs | hypothesis). The likelihood is a distribution that is proportional to the probability of the observed data given the hypothesis.12 Each parameter λi and τ of our prior probability distribution has its own likelihood function. In other words, the different values of λi have likelihood functions that are normal random variables with means equal to the observed means (here, 7 ant nests per quadrat in the forest and 10.75 ant nests per quadrat in the field; see Table 5.2). The variances are equal to the sample variances (4.79 in the forest and 2.25 in the field). The parameter τ has a likelihood function that is a gamma random variable. Finally, the F-ratio is an F-random variable with expected value (or maximum likelihood estimate)13 equal to 8.78 (calculated from the data using Equation 5.1). Step 5: Calculating the Posterior Probability Distribution

To calculate the posterior probability distribution, P(H | data), we apply Equation 5.2, multiply the prior by the likelihood, and divide by the normalizing constant (or marginal likelihood). Although this multiplication is straightforward for well-behaved distributions like the normal, computational methods are used

12 There is a key difference between the likelihood function and a probability distribution. The probability of data given a hypothesis, P(data | H), is the probability of any set of random data given a specific hypothesis, usually the statistical null hypothesis. The associated probability density function (see Chapter 2) conforms to the First Axiom of Probability—that the sum of all probabilities = 1. In contrast, the likelihood is based on only one dataset (the observed sample) and may be calculated for many different hypotheses or parameters. Although it is a function, and results in a distribution of values, the distribution is not a probability distribution, and the sum of all likelihoods does not necessarily sum to 1. 13

The maximum likelihood is the value for our parameter that maximizes the likelihood function. To obtain this value, take the derivative of the likelihood, set it to 0, and solve for the parameter values. Frequentist parameter estimates are usually equal to maximum-likelihood estimates for the parameters of the specified probability density functions. Fisher claimed, in his system of fiducial inference, that the maximum-likelihood estimate gave a realistic probability of the alternative (or null) hypothesis. Fisher based this claim on a statistical axiom that he defined by saying that given observed data Y, the likelihood function L(H | Y) contains all the relevant information about the hypothesis H. See Chapter 14 for further application of maximum likelihood and Bayesian methods. Berger and Wolpert (1984), Edwards (1992), and Bolker (2008) provide additional discussion of likelihood methods.

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to iteratively estimate the posterior distribution for any prior distribution (Carlin and Louis 2000; Kéry 2010). In contrast to the results of a parametric or Monte Carlo analysis, the result of a Bayesian analysis is a probability distribution, not a single P-value. Thus, in this example, we express P(F 5.32 | Fobs) as a random variable with expected mean and variance. For the data in Table 5.1, we calculated posterior estimates for all the parameters: λfield, λforest, σ2 (= 1/τ) (Table 5.7). Because we used uninformative priors, the parameter estimates for the Bayesian and parametric analyses are similar, though not identical. The hypothesized F-distribution with expected value equal to 5.32 is shown in Figure 5.8. To compute P(F ≥ 5.32 | Fobs), we simulated 20,000 F-ratios using a Monte Carlo algorithm. The average, or expected value, of all of these F-ratios is 9.77; this number is somewhat larger than the frequentist (maximum likelihood) estimate of 8.78 because our sample size is very small (N = 10). The spread about the mean is large: SD = 7.495; hence the precision of our estimate is relatively low (0.017). Step 6: Interpreting the Results

We now return to the motivating question: What is the probability of obtaining an F-ratio ≥ 5.32, given the data on ant nest density in Table 5.1? In other words, how probable is it that the mean ant nest densities in the two habitats really differ? We can answer this question directly by asking what percentage of values in Figure 5.8 are greater than or equal to 5.32. The answer is 67.3%. This doesn’t look quite as convincing as the P-value of 0.018 (1.8%) obtained in the parametric analysis in the previous section. In fact, the percentage of values in Figure 5.8 that are ≥8.78, the observed value for which we found P = 0.018 in the parametric analysis section, is 46.5. In other words, the Bayesian analysis (see Figure 5.8) TABLE 5.7 Parametric and Bayesian estimators for the means and standard deviations of the data in Table 5.1 Estimator Analysis

lForest

lField

sForest

sField

Parametric Bayesian (uninformed prior) Bayesian (informed prior)

7.00 6.97 7.00

10.75 10.74 10.74

2.19 0.91 1.01

1.50 1.13 1.02

The standard deviation estimators from the Bayesian analysis are slightly smaller because the Bayesian analysis incorporates information from the prior probability distribution. Bayesian analysis may give different results, depending on the shape of the prior distribution and the sample size.

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Hypothetical value of F = 5.32

Probability

0.06

Observed value of F = 9.77

0.04

0.02

0.00 0

30 F-ratio

Figure 5.8 Hypothesized F-distribution with an expected value of 5.32. We are interested in determining the probability of F ≥ 5.32 (the critical value for P < 0.05 in a standard F-ratio test), given the data on ant nest densities in Table 5.1. This is the inverse of the traditional null hypothesis, which asks: what is the probability of obtaining the data, given the null hypothesis? In the Bayesian analysis, the posterior probability of F ≥ 5.32, given the data in Table 5.1, is the proportion of the area under the curve to the right of F = 5.32, which is 0.673. In other words, P(hypothesis that the fields and forests differ in average density of ant nests | observed data in Table 5.1) = 0.673. The most likely posterior value of the F-ratio is 9.77. The proportion of area under the curve to the right of this value is 0.413. The parametric analysis says that the observed data are unlikely given a null distribution specified by the F-ratio [P(data | H0) = 0.018], whereas the Bayesian analysis says that the probability of observing an F-ratio of 9.77 or larger is not unlikely given the data [P(F 5.32 | data) = 0.673].

indicates P = 0.67 that ant nest densities in the two habitats are truly different, given the Bayesian estimate of F = 9.77 [P(F 5.32 | Fobs) = 0.67]. In contrast, the parametric analysis (see Figure 5.5) indicates P = 0.018 that the parametric estimate of F = 8.78 (or a greater F-ratio) would be found given the null hypothesis that the ant densities in the two habitats are the same [P(Fobs | H0) = 0.018]. Using Bayesian analysis, a different answer would result if we used a different prior distribution rather than the uninformative prior of means of 0 with large variances. For example, if we used prior means of 15 for the forest and 7 for the field, an among-group variance of 10, and a within-group variance of 0.001, then P(F ≥ 5.32 | data) = 0.57. Nevertheless, you can see that the posterior probability does depend on the priors that are used in the analysis (see Table 5.7). Finally, we can estimate a 95% Bayesian credibility interval around our estimate of the observed F-ratio. As with Monte Carlo methods, we estimate the

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95% credibility interval as the 2.5 and 97.5 percentiles of the simulated F-ratios. These values are 0.28 and 28.39. Thus, we can say that we are 95% sure that the value of the F-ratio for this experiment lies in the interval [0.28, 28.39]. Note that the spread is large because the precision of our estimate of the F-ratio is low, reflecting the small sample size in our analysis. You should compare this interpretation of a credibility interval with the interpretation of a confidence interval presented in Chapter 3. Assumptions of Bayesian Analysis

In addition to the standard assumptions of all statistics methods (random, independent observations), the key assumption of Bayesian analysis is that the parameters to be estimated are random variables with known distributions. In our analysis, we also assumed little prior information (uninformative priors), and therefore the likelihood function had more influence on the final calculation of the posterior probability distribution than did the prior. This should make intuitive sense. On the other hand, if we had a lot of prior information, our prior probability distribution (e.g., the black curve in Figure 5.6) would have low variance and the likelihood function would not substantially change the variance of the posterior probability distribution. If we had a lot of prior information, and we were confident in it, we would not have learned much from the experiment. A well-designed experiment should decrease the posterior estimate of the variance relative to the prior estimate of the variance. The relative contributions of prior and likelihood to the posterior estimate of the probability of the mean density of nests in the forest are illustrated in Figure 5.9. In this figure, the prior is flat over the range of the data (i.e., it is an unin-

0.04

Figure 5.9 Probability densities for the prior, likelihood,

0.03 Probability

and posterior for the mean number of ant nests in the forest plots. In the Bayesian analysis of the data in Table 5.1, we used an uninformative prior distribution with a mean of 0 and a variance of 100,000. This normal distribution generated an essentially uniform range of prior probability values (dark blue) over the range of values for λforest, the density of forest ants. The likelihood (light blue) represents the probability based on the observed data (see Table 5.1), and the posterior probability (black) is the product of the two. Notice that the posterior distribution is more precise than the likelihood because it takes into account the (modest) information contained in the prior.

0.02

0.01

0.00 0.0

2.5

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7.5

λ forest

10.0

12.5

Summary

formative prior), the likelihood is the distribution based on the observed data (see Table 5.1), and the posterior is the product of the two. Note that the variance of the posterior is smaller than the variance of the likelihood because we had some prior information. However, the expected values of the likelihood (7.0) and the posterior (6.97) are very close, because all values of the prior were approximately equally likely over the range of the data. After doing this experiment, we have new information on each of the parameters that could be used in analysis of subsequent experiments. For example, if we were to repeat the experiment, we could use the posterior in Figure 5.9 as our prior for the average density of ant nests in other forests. To do this, we would use the values for λi and σi in Table 5.7 as the estimates of λi and σi in setting up the new prior probability distributions. Advantages and Disadvantages of Bayesian Analysis

Bayesian analysis has a number of advantages relative to parametric and Monte Carlo approaches conducted in a frequentist framework. Bayesian analysis allows for the explicit incorporation of prior information, and the results from one experiment (the posterior) can be used to inform (as a prior) subsequent experiments. The results of Bayesian analysis are interpreted in an intuitively straightforward way, and the inferences obtained are conditional on both the observed data and the prior information. Disadvantages to Bayesian analysis are its computational challenges (Albert 2007; Clark 2007; Kéry 2007) and the requirement to condition the hypothesis on the data [i.e., P(hypothesis | data)]. The most serious disadvantage of Bayesian analysis is its potential lack of objectivity, because different results will be obtained using different priors. Consequently, different investigators may obtain different results from the same dataset if they start with different preconceptions or prior information. The use of uninformative priors addresses this criticism, but increases the computational complexity.

Summary Three major frameworks for statistical analysis are Monte Carlo, parametric, and Bayesian. All three assume that the data were sampled randomly and independently. In Monte Carlo analysis, the data are randomized or reshuffled, so that individuals are randomly re-assigned to groups. Test statistics are calculated for these randomized datasets, and the reshuffling is repeated many times to generate a distribution of simulated values. The tail probability of the observed test statistic is then estimated from this distribution. The advantage of Monte Carlo analysis is that it makes no assumptions about the distribution of the data,

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it makes the null hypothesis clear and explicit, and it can be tailored to individual datasets and hypotheses. The disadvantage is that it is not a general solution and usually requires computer programming to be implemented. In parametric analysis, the data are assumed to have been drawn from an underlying known distribution. An observed test statistic is compared to a theoretical distribution based on a null hypothesis of random variation. The advantage of parametric analysis is that it provides a unifying framework for statistical tests of classical null hypotheses. Parametric analysis is also familiar to most ecologists and environmental scientists and is widely implemented in statistical software. The disadvantage of parametric analysis is that the tests do not specify the probability of alternative hypotheses, which often is of greater interest than the null hypothesis. Bayesian analysis considers parameters to be random variables as opposed to having fixed values. It can take explicit advantage of prior information, although modern Bayesian methods rely on uninformative priors. The results of Bayesian analysis are expressed as probability distributions, and their interpretation conforms to our intuition. However, Bayesian analysis requires complex computation and often requires the investigators to write their own programs.

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PART II

Designing Experiments

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Designing Successful Field Studies

The proper analysis of data goes hand in hand with an appropriate sampling design and experimental layout. If there are serious errors or problems in the design of the study or in the collection of the data, rarely is it possible to repair these problems after the fact. In contrast, if the study is properly designed and executed, the data can often be analyzed in several different ways to answer different questions. In this chapter, we discuss the broad issues that you need to consider when designing an ecological study. We can’t overemphasize the importance of thinking about these issues before you begin to collect data.

What Is the Point of the Study? Although it may seem facetious and the answer self-evident, many studies are initiated without a clear answer to this central question. Most answers will take the form of a more focused question. Are There Spatial or Temporal Differences in Variable Y?

This is the most common question that is addressed with survey data, and it represents the starting point of many ecological studies. Standard statistical methods such as analysis of variance (ANOVA) and regression are well-suited to answer this question. Moreover, the conventional testing and rejection of a simple null hypothesis (see Chapter 4) yields a dichotomous yes/no answer to this question. It is difficult to even discuss mechanisms without some sense of the spatial or temporal pattern in your data. Understanding the forces controlling biological diversity, for example, requires at a minimum a spatial map of species richness. The design and implementation of a successful ecological survey requires a great deal of effort and care, just as much as is needed for a successful experimental study. In some cases, the survey study will address all of your research goals; in other cases, a survey study will be the first step in a research

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project. Once you have documented spatial and temporal patterns in your data, you will conduct experiments or collect additional data to address the mechanisms responsible for those patterns. What Is the Effect of Factor X on Variable Y?

This is the question directly answered by a manipulative experiment. In a field or laboratory experiment, the investigator actively establishes different levels of Factor X and measures the response of Variable Y. If the experimental design and statistical analysis are appropriate, the resulting P-value can be used to test the null hypothesis of no effect of Factor X. Statistically significant results suggest that Factor X influences Variable Y, and that the “signal” of Factor X is strong enough to be detected above the “noise” caused by other sources of natural variation.1 Certain natural experiments can be analyzed in the same way, taking advantage of natural variation that exists in Factor X. However, the resulting inferences are usually weaker because there is less control over confounding variables. We discuss natural experiments in more detail later in this chapter. Are the Measurements of Variable Y Consistent with the Predictions of Hypothesis H?

This question represents the classic confrontation between theory and data (Hilborn and Mangel 1997). In Chapter 4, we discussed two strategies we use for this confrontation: the inductive approach, in which a single hypothesis is recursively modified to conform to accumulating data, and the hypotheticodeductive approach, in which hypotheses are falsified and discarded if they do not predict the data. Data from either experimental or observational studies can be used to ask whether observations are consistent with the predictions of a mechanistic hypothesis. Unfortunately, ecologists do not always state this question so plainly. Two limitations are (1) many ecological hypotheses do not generate simple, falsifiable predictions; and (2) even when an hypothesis does generate predictions, they are rarely unique. Therefore, it may not be possible to definitively test Hypothesis H using only data collected on Variable Y.

Although manipulative experiments allow for strong inferences, they may not reveal explicit mechanisms. Many ecological experiments are simple “black box” experiments that measure the response of the Variable Y to changes in Factor X, but do not elucidate lower-level mechanisms causing that response. Such a mechanistic understanding may require additional observations or experiments addressing a more focused question about process.

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Manipulative Experiments

Using the Measurements of Variable Y, What Is the Best Estimate of Parameter q in Model Z?

Statistical and mathematical models are powerful tools in ecology and environmental science. They allow us to forecast how populations and communities will change through time or respond to altered environmental conditions (e.g., Sjögren-Gulve and Ebenhard 2000). Models can also help us to understand how different ecological mechanisms interact simultaneously to control the structure of communities and populations (Caswell 1988). Parameter estimation is required for building predictive models and is an especially important feature of Bayesian analysis (see Chapter 5). Rarely is there a simple one-toone correspondence between the value of Variable Y measured in the field and the value of Parameter θ in our model. Instead, those parameters have to be extracted and estimated indirectly from our data. Unfortunately, some of the most common and traditional designs used in ecological experiments and field surveys, such as the analysis of variance (see Chapter 10), are not very useful for estimating model parameters. Chapter 7 discusses some alternative designs that are more useful for parameter estimation.

Manipulative Experiments In a manipulative experiment, the investigator first alters levels of the predictor variable (or factor), and then measures how one or more variables of interest respond to these alterations. These results are then used to test hypotheses of cause and effect. For example, if we are interested in testing the hypothesis that lizard predation controls spider density on small Caribbean islands, we could alter the density of lizards in a series of enclosures and measure the resulting density of spiders (e.g., Spiller and Schoener 1998). We could then plot these data in a graph in which the x-axis (= independent variable) is lizard density, and the y-axis (= dependent variable) is spider density (Figure 6.1A,B). Our null hypothesis is that there is no relationship between these two variables (Figure 6.1A). That is, spider density might be high or low in a particular enclosure, but it is not related to the density of lizards that were established in the enclosure. Alternatively, we might observe a negative relationship between spider and lizard density: enclosures with the highest lizard density have the fewest spiders, and vice-versa (Figure 6.1B). This pattern then would have to be subject to a statistical analysis such as regression (see Chapter 9) to determine whether or not the evidence was sufficient to reject the null hypothesis of no relationship between lizard and spider densities. From these data we could also estimate regression model parameters that quantify the strength of the relationship.

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(A) Null hypothesis

(B) Alternative hypothesis

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Figure 6.1 Relationship between lizard density and spider density in manipulative and natural field experiments. Each point represents a plot or quadrat in which both spider density and lizard density have been measured. (A) The null hypothesis is that lizard density has no effect on spider density. (B) The alternative hypothesis is that lizard predation controls spider density, leading to a negative relationship between these two variables.

Although field experiments are popular and powerful, they have several important limitations. First, it is challenging to conduct experiments on large spatial scales; over 80% of field experiments have been conducted in plots of less than 1 m2 (Kareiva and Anderson 1988; Wiens 1989). When experiments are conducted on large spatial scales, replication is inevitably sacrificed (Carpenter 1989). Even when they are properly replicated, experiments conducted on small spatial scales may not yield results that are representative of patterns and processes occurring at larger spatial scales (Englund and Cooper 2003). Second, field experiments are often restricted to relatively small-bodied and short-lived organisms that are easy to manipulate. Although we always want to generalize the results of our experiments to other systems, it is unlikely that the interaction between lizards and spiders will tell us much about the interaction between lions and wildebeest. Third, it is difficult to change one and only one variable at a time in a manipulative experiment. For example, cages can exclude other kinds of predators and prey, and introduce shading. If we carelessly compare spider densities in caged plots versus uncaged “controls,” the effects of lizard predation are confounded with other physical differences among the treatments. We discuss solutions to confounding variables later in this chapter. Finally, many standard experimental designs are simply unwieldy for realistic field experiments. For example, suppose we are interested in investigating competitive interactions in a group of eight spider species. Each treatment in

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Natural Experiments

such an experiment would consist of a unique combination of species. Although the number of species in each treatment ranges from only 1 to 8, the number of unique combinations is 28 – 1 = 255. If we want to establish even 10 replicates of each treatment (see “The Rule of 10,” discussed later in this chapter), we need 2550 plots. That may not be possible because of constraints on space, time, or labor. Because of all these potential limitations, many important questions in community ecology cannot be addressed with field experiments.

Natural Experiments A natural experiment (Cody 1974) is not really an experiment at all. Instead, it is an observational study in which we take advantage of natural variation that is present in the variable of interest. For example, rather than manipulate lizard densities directly (a difficult, expensive, and time-consuming endeavor), we could census a set of plots (or islands) that vary naturally in their density of lizards (Schoener 1991). Ideally, these plots would vary only in the density of lizards and would be identical in all other ways. We could then analyze the relationship between spider density and lizard density as illustrated in Figure 6.1. Natural experiments and manipulative experiments superficially generate the same kinds of data and are often analyzed with the same kinds of statistics. However, there are often important differences in the interpretation of natural and manipulative experiments. In a manipulative experiment, if we have established valid controls and maintained the same environmental conditions among the replicates, any consistent differences in the response variable (e.g., spider density) can be attributed confidently to differences in the manipulated factor (e.g., lizard density). We don’t have this same confidence in interpreting results of natural experiments. In a natural experiment, we do not know the direction of cause and effect, and we have not controlled for other variables that surely will differ among the replicates. For the lizard–spider example, there are at least four hypotheses that could account for a negative association between lizard and spider densities: 1. Lizards may control spider density. This was the alternative hypothesis of interest in the original field experiment. 2. Spiders may directly or indirectly control lizard density. Suppose, for example, that large hunting spiders consume small lizards, or that spiders are also preyed upon by birds that feed on lizards. In both cases, increasing spider density may decrease lizard density, even though lizards do feed on spiders.

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3. Both spider and lizard densities are controlled by an unmeasured environmental factor. For example, suppose that spider densities are highest in wet plots and lizard densities are highest in dry plots. Even if lizards have little effect on spiders, the pattern in Figure 6.1B will emerge: wet plots will have many spiders and few lizards, and dry plots will have many lizards and few spiders. 4. Environmental factors may control the strength of the interaction between lizards and spiders. For example, lizards might be efficient predators on spiders in dry plots, but inefficient predators in wet plots. In such cases, the density of spiders will depend on both the density of lizards and the level of moisture in the plot (Spiller and Schoener 1995). These four scenarios are only the simplest ones that might lead to a negative relationship between lizard density and spider density (Figure 6.2). If we add double-headed arrows to these diagrams (lizards and spiders reciprocally affect one another’s densities), there is an even larger suite of hypotheses that could account for the observed relationships between spider density and lizard density (see Figure 6.1). All of this does not mean that natural experiments are hopeless, however. In many cases we can collect additional data to distinguish among these hypotheses. For example, if we suspect that environmental variables such as moisture

Lizard density

Spider density

Lizard density

Moisture

Lizard density

Spider density

Moisture

Spider density

Lizard density

Spider density

Figure 6.2 Mechanistic hypotheses to account for correlations between lizard density and spider density (see Figure 6.1). The cause-and-effect relationship might be from predator to prey (upper left) or prey to predator (upper right). More complicated models include the effects of other biotic or abiotic variables. For example, there might be no interaction between spiders and lizards, but densities of both are controlled by a third variable, such as moisture (lower left). Alternatively, moisture might have an indirect effect by altering the interaction of lizards and spiders (lower right).

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Snapshot versus Trajectory Experiments

are important, we either can restrict the survey to a set of plots with comparable moisture levels, or (better still) measure lizard density, spider density, and moisture levels in a series of plots censused over a moisture gradient. Confounding variables and alternative mechanisms also can be problematic in field experiments. However, their impacts will be reduced if the investigator conducts the experiment at an appropriate spatial and temporal scale, establishes proper controls, replicates adequately, and uses randomization to locate replicates and assign treatments. Overall, manipulative experiments allow for greater confidence in our inferences about cause and effect, but they are limited to relatively small spatial scales and short time frames. Natural experiments can be conducted at virtually any spatial scale (small quadrats to entire continents) and over any time interval (weekly field measurements, to annual censuses, to fossil strata). However, it is more challenging to tease apart cause-and-effect relationships in natural experiments.2

Snapshot versus Trajectory Experiments Two variants of the natural experiment are the snapshot experiment and the trajectory experiment (Diamond 1986). Snapshot experiments are replicated in space, and trajectory experiments are replicated in time. For the data in Figure 6.1, suppose we censused 10 different plots in a single day. This is a snapshot experiment in which the replication is spatial; each observation represents a different plot censused at the same time. On the other hand, suppose we visited a single plot in 10 different years. This is a trajectory experiment in which the replication is temporal; each observation represents a different year in the study. The advantages of a snapshot experiment are that it is rapid, and the spatial replicates arguably are more statistically independent of one another than are

In some cases, the distinction between manipulative and natural field experiments is not clear-cut. Human activity has generated many unintended large-scale experiments including eutrophication, habitat alteration, global climate change, and species introductions and removals. Imaginative ecologists can take advantage of these alterations to design studies in which the confidence in the conclusions is very high. For example, Knapp et al. (2001) studied the impacts of trout introductions to lakes in the Sierra Nevada by comparing invertebrate communities in naturally fishless lakes, stocked lakes, and lakes that formerly were stocked with fish. Many comparisons of this kind are possible to document consequences of human activity. However, as human impacts become more widespread and pervasive, it may be harder and harder to find sites that can be considered unmanipulated controls.

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the temporal replicates of a trajectory experiment. The majority of ecological data sets are snapshot experiments, reflecting the 3- to 5-year time frame of most research grants and dissertation studies.3 In fact, many studies of temporal change are actually snapshot studies, because variation in space is treated as a proxy variable for variation in time. For example, successional change in plant communities can be studied by sampling from a chronosequence—a set of observations, sites, or habitats along a spatial gradient that differ in the time of origin (e.g., Law et al. 2003). The advantage of a trajectory experiment is that it reveals how ecological systems change through time. Many ecological and environmental models describe precisely this kind of change, and trajectory experiments allow for stronger comparisons between model predictions and field data. Moreover, many models for conservation and environmental forecasting are designed to predict future conditions, and data for these models are derived most reliably from trajectory experiments. Many of the most valuable data sets in ecology are long time-series data for which populations and communities at a site are sampled year after year with consistent, standardized methods. However, trajectory experiments that are restricted to a single site are unreplicated in space. We don’t know if the temporal trajectories described from that site are typical for what we might find at other sites. Each trajectory is essentially a sample size of one at a given site.4 The Problem of Temporal Dependence

A more difficult problem with trajectory experiments is the potential non-independence of data collected in a temporal sequence. For example, suppose you measure tree diameters each month for one year in a plot of redwood trees. Red-

A notable exception to short-term ecological experiments is the coordinated set of studies developed at Long Term Ecological Research (LTER) sites. The National Science Foundation (NSF) funded the establishment of these sites throughout the 1980s and 1990s specifically to address the need for ecological research studies that span decades to centuries. See www.lternet.edu/.

Snapshot and trajectory designs show up in manipulative experiments as well. In particular, some designs include a series of measurements taken before and after a manipulation. The “before” measurements serve as a type of “control” that can be compared to the measurements taken after the manipulation or intervention. This sort of BACI design (Before-After, Control-Impact) is especially important in environmental impact analysis and in studies where spatial replication may be limited. For more on BACI, see the section “Large Scale Studies and Environmental Impacts” later in this chapter, and see Chapter 7.

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Snapshot versus Trajectory Experiments

woods grow very slowly, so the measurements from one month to the next will be virtually identical. Most foresters would say that you don’t have 12 independent data points, you have only one (the average diameter for that year). On the other hand, monthly measurements of a rapidly developing freshwater plankton community reasonably could be viewed as statistically independent of one another. Naturally, the further apart in time the samples are separated from one another, the more they function as independent replicates. But even if the correct census interval is used, there is still a subtle problem in how temporal change should be modeled. For example, suppose you are trying to model changes in population size of a desert annual plant for which you have access to a nice trajectory study, with 100 years of consecutive annual censuses. You could fit a standard linear regression model (see Chapter 9) to the time series Nt = β0 + β1t + ε

(6.1)

In this equation, population size (Nt) is a linear function of the amount of time (t) that has passed. The coefficients β0 and β1 are the intercept and slope of this straight line. If β1 is less than 0.0, the population is shrinking with time, and if β1 > 0, N is increasing. Here ε is a normally distributed white noise5 error term that incorporates both measurement error and random variation in population size. Chapter 9 will explain this model in much greater detail, but we introduce it now as a simple way to think about how population size might change in a linear fashion with the passage of time. However, this model does not take into account that population size changes through births and deaths affecting current population size. A time-series model would describe population growth as Nt+1 = β0 + β1Nt + ε

(6.2)

White noise is a type of error distribution in which the errors are independent and uncorrelated with one another. It is called white noise as an analogy to white light, which is an equal mixture of short and long wavelengths. In contrast, red noise is dominated by low-frequency perturbations, just as red light is dominated by low-frequency light waves. Most time series of population sizes exhibit a reddened noise spectrum (Pimm and Redfearn 1988), so that variances in population size increase when they are analyzed at larger temporal scales. Parametric regression models require normally distributed error terms, so white noise distributions form the basis for most stochastic ecological models. However, an entire spectrum of colored noise distributions (1/f noise) may provide a better fit to many ecological and evolutionary datasets (Halley 1996).

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In this model, the population size in the next time step (Nt+1) depends not simply on the amount of time t that has passed, but rather on the population size at the last time step (Nt). In this model, the constant β1 is a multiplier term that determines whether the population is exponentially increasing (β1 > 1.0) or decreasing (β1 < 1.0). As before, ε is a white noise error term. The linear model (Equation 6.1) describes a simple additive increase of N with time, whereas the time-series, or autoregressive model (Equation 6.2) describes an exponential increase, because the factor β1 is a multiplier that, on average, gives a constant percentage increase in population size at each time step. The more important difference between the two models, however, is that the differences between the observed and predicted population sizes (i.e., the deviations) in the time-series model are correlated with one another. As a consequence, there tend to be runs, or periods of consecutive increases followed by periods of consecutive decreases. This is because the growth trajectory has a “memory”—each consecutive observation (Nt+1) depends directly on the one that came before it (the Nt term in Equation 6.2). In contrast, the linear model has no memory, and the increases are a function only of time (and ε), and not of Nt. Hence, the positive and negative deviations follow one another in a purely random fashion (Figure 6.3). Correlated deviations, which are typical of data collected in trajectory studies, violate the assumptions of most conventional statistical analyses.6 Analytical and computer-intensive methods have been developed for analyzing both sample data and experimental data collected through time (Ives et al. 2003; Turchin 2003). This does not mean we cannot incorporate time-series data into conventional statistical analyses. In Chapters 7 and 10, we will discuss additional ways to analyze time-series data. These methods require that you pay careful attention to both the sampling design and the treatment of the data after you have collected them. In this respect, time-series or trajectory data are just like any other data.

Press versus Pulse Experiments In manipulative studies, we also distinguish between press experiments and pulse experiments (Bender et al. 1984). In a press experiment, the altered conditions in the treatment are maintained through time and are re-applied as necessary to ensure that the strength of the manipulation remains constant. Thus, 6

Actually, spatial autocorrelation generates the same problems (Legendre and Legendre 1998; Lichstein et al. 2003). However, tools for spatial autocorrelation analysis have developed more or less independently of time-series analyses, perhaps because we perceive time as a strictly one-dimensional variable and space as a two- or threedimensional variable.

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105 100 95

90 0

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Time

Figure 6.3 Examples of deterministic and stochastic time series, with and without autocorrelation. Each population begins with 100 individuals. A linear model without error (dashed line) illustrates a constant upward trend in population data. A linear model with a stochastic white noise error term (black line) adds temporally uncorrelated variability. Finally, an autocorrelated model (blue line) describes population size in the next time step (t + 1) as a function of the population size in the current time step (t) plus random noise. Although the error term in this model is still a simple random variable, the resulting time series shows autocorrelation— there are runs of population increases followed by runs of population decreases. For the linear model and the stochastic white noise model, the equation is Nt = a + bt + ε, with a = 100 and b = 0.10. For the autocorrelated model, Nt+1 = a + bNt + ε, with a = 0.0 and b = 1.0015. For both models with error, ε is a normal random variable: ε ~ N(0,1).

fertilizer may have to be re-applied to plants, and animals that have died or disappeared from a plot may have to be replaced. In contrast, in a pulse experiment, experimental treatments are applied only once, at the start of the study. The treatment is not re-applied, and the replicate is allowed to “recover” from the manipulation (Figure 6.4A). Press and pulse experiments measure two different responses to the treatment. The press experiment (Figure 6.4B) measures the resistance of the system to the experimental treatment: the extent to which it resists change in the constant environment created by the press experiment. A system with low resistance will exhibit a large response in a press experiment, whereas a system with high resistance will exhibit little difference between control and manipulated treatments. The pulse experiment measures the resilience of the system to the experimental treatment: the extent to which the system recovers from a single perturbation. A system with high resilience will show a rapid return to control con-

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(A) Pulse experiment

(B) Press experiment

Response variable

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Time

Figure 6.4 Ecological pulse and press experiments. The arrow indicates a treatment application, and the line indicates the temporal trajectory of the response variable. The pulse experiment (A) measures the response to a single treatment application (resilience), whereas the press experiment (B) measures the response under constant conditions (resistance).

ditions, whereas a system with low resilience will take a long time to recover; control and manipulated plots will continue to differ for a long time after the single treatment application. The distinction between press and pulse experiments is not in the number of treatment applications used, but in whether the altered conditions are maintained through time in the treatment. If environmental conditions remain constant following a single perturbation for the duration of the experiment, the design is effectively a press experiment. Another distinction between press and pulse experiments is that the press experiment measures the response of the system under equilibrium conditions, whereas the pulse experiment records transient responses in a changing environment.

Replication How Much Replication?

This is one of the most common questions that ecologists and environmental scientists ask of statisticians. The correct response is that the answer depends on the variance in the data and the effect size—the difference that you wish to detect between the averages of the groups being compared. Unfortunately, these two quantities may be difficult to estimate, although you always should consider what effect size would be reasonable to observe. To estimate variances, many statisticians will recommend that you conduct a pilot study. Unfortunately, pilot studies usually are not feasible—you rarely have the freedom to set up and run a costly or lengthy study more than once.

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Replication

Field seasons and grant proposals are too short for this sort of luxury. However, you may be able to estimate reasonable ranges of variances and effect sizes from previously published studies and from discussions with colleagues. You can then use these values to determine the statistical power (see Chapter 4) that will result from different combinations of replicates, variances, and effect sizes (see Figure 4.5 for an example). At a minimum, however, you need to first answer the following question: How Many Total Replicates Are Affordable?

It takes time, labor, and money to collect either experimental or survey data, and you need to determine precisely the total sample size that you can afford. If you are conducting expensive tissue or sample analyses, the dollar cost may be the limiting factor. However, in many studies, time and labor are more limiting than money. This is especially true for geographical surveys conducted over large spatial scales, for which you (and your field crew if you are lucky enough to have one) may spend as much time traveling to study sites as you do collecting field data. Ideally, all of the replicates should be measured simultaneously, giving you a perfect snapshot experiment. The more time it takes to collect all the data, the more conditions will have changed from the first sample to the last. For experimental studies, if the data are not collected all at once, then the amount of time that has passed since treatment application is no longer identical for all replicates. Obviously, the larger the spatial scale of the study, the harder it is to collect all of the data within a reasonable time frame. Nevertheless, the payoff may be greater because the scope of inference is tied to the spatial scale of analysis: conclusions based on samples taken only at one site may not be valid at other sites. However, there is no point in developing an unrealistic sampling design. Carefully map out your project from start to finish to ensure it will be feasible.7 Only once you know the total number of replicates or observations that you can collect can you begin to design your experiment by applying the rule of 10.

It can be very informative to use a stopwatch to time carefully how long it takes to complete a single replicate measurement of your study. Like the efficiency expert father in Cheaper By The Dozen (Gilbreth and Carey 1949), we put great stock in such numbers. With these data, we can accurately estimate how many replicates we can take in an hour, and how much total field time we will need to complete the census. The same principle applies to sample processing, measurements that we make back in the laboratory, the entry of data into the computer, and the long-term storage and curation of data (see Chapter 8). All of these activities take time that needs to be accounted for when planning an ecological study.

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The Rule of 10

The Rule of 10 is that you should collect at least 10 replicate observations for each category or treatment level. For example, suppose you have determined that you can collect 50 total observations in a experiment examining photosynthetic rates among different plant species. A good design for a one-way ANOVA would be to compare photosynthetic rates among not more than five species. For each species, you would choose randomly 10 plants and take one measurement from each plant. The Rule of 10 is not based on any theoretical principle of experimental design or statistical analysis, but is a reflection of our hard-won field experience with designs that have been successful and those that have not. It is certainly possible to analyze data sets with less than 10 observations per treatment, and we ourselves often break the rule. Balanced designs with many treatment combinations but only four or five replicates may be quite powerful. And certain one-way designs with only a few treatment levels may require more than 10 replicates per treatment if variances are large. Nevertheless, the Rule of 10 is a solid starting point. Even if you set up the design with 10 observations per treatment level, it is unlikely that you will end up with that number. In spite of your best efforts, data may be lost for a variety of reasons, including equipment failures, weather disasters, plot losses, human disturbances or errors, improper data transcription, and environmental alterations. The Rule of 10 at least gives you a fighting chance to collect data with reasonable statistical power for revealing patterns.8 In Chapter 7, we will discuss efficient sample designs and strategies for maximizing the amount of information you can squeeze out of your data. Large-Scale Studies and Environmental Impacts

The Rule of 10 is useful for small-scale manipulative studies in which the study units (plots, leaves, etc.) are of manageable size. But it doesn’t apply to large-scale ecosystem experiments, such as whole-lake manipulations, because replicates may be unavailable or too expensive. The Rule of 10 also does not apply to many environmental impact studies, where the assessment of an impact is required at a single site. In such cases, the best strategy is to use a BACI design (Before-After, Control-Impact). In some BACI designs, the replication is achieved through time:

Another useful rule is the Rule of 5. If you want to estimate the curvature or non-linearity of a response, you need to use at least five levels of the predictor variable. As we will discuss in Chapter 7, a better solution is to use a regression design, in which the predictor variable is continuous, rather than categorical with a fixed number of levels.

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Ensuring Independence

the control and impact sites are censused repeatedly both before and after the impact. The lack of spatial replication restricts the inferences to the impact site itself (which may be the point of the study), and requires that the impact is not confounded with other factors that may be co-varying with the impact. The lack of spatial replication in simple BACI designs is controversial (Underwood 1994; Murtaugh 2002b), but in many cases they are the best design option (Stewart-Oaten and Bence 2001), especially if they are used with explicit time-series modeling (Carpenter et al. 1989). We will return to BACI and its alternatives in Chapters 7 and 10.

Ensuring Independence Most statistical analyses assume that replicates are independent of one another. By independence, we mean that the observations collected in one replicate do not have an influence on the observations collected in another replicate. Nonindependence is most easily understood in an experimental context. Suppose you are studying the response of hummingbird pollinators to the amount of nectar produced by flowers. You set up two adjacent 5 m × 5 m plots. One plot is a control plot; the adjacent plot is a nectar removal plot in which you drain all of the nectar from the flowers. You measure hummingbird visits to flowers in the two plots. In the control plot, you measure an average of 10 visits/hour, compared to only 5 visits/hour in the removal plot. However, while collecting the data, you notice that once birds arrive at the removal plot, they immediately leave, and the same birds then visit the adjacent control plot (Figure 6.5A). Clearly, the two sets of observations are not independent of one another. If the control and treatment plots had been more widely separated in space, the numbers might have come out differently, and the average in the control plots might have been only 7 visits/hour instead of 10 visits/hour (Figure 6.5B). When the two plots are adjacent to one another, non-independence inflates the difference between them, perhaps leading to a spuriously low P-value, and a Type I error (incorrect rejection of a true null hypothesis; see Chapter 4). In other cases, non-independence may decrease the apparent differences between treatments, contributing to a Type II error (incorrect acceptance of a false null hypothesis). Unfortunately, non-independence inflates or deflates both P-values and power to unknown degrees. The best safeguard against non-independence is to ensure that replicates within and among treatments are separated from one another by enough space or time to ensure that the they do not affect one another. Unfortunately, we rarely know what that distance or spacing should be, and this is true for both experimental and observational studies. We should use common sense and as much

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Figure 6.5 The problem of non-independence in ecological studies is illustrated by an experimental design in which hummingbirds forage for nectar in control plots and in plots from which nectar has been removed from all of the flowers. (A) In a non-independent layout, the nectar removal and control plots are adjacent to one another, and hummingbirds that enter the nectar removal plot immediately leave and begin foraging in the adjacent control plot. As a consequence, the data collected in the control plot are not independent of the data collected in the nectar removal plot: the responses in one treatment influence the responses in the other. (B) If the layout is modified so that the two plots are widely separated, hummingbirds that leave the nectar removal plot do not necessarily enter the control plot. The two plots are independent, and the data collected in one plot are not influenced by the presence of the other plot. Although it is easy to illustrate the potential problem of nonindependence, in practice it is can be very difficult to know ahead of time the spatial and temporal scales that will ensure statistical independence.

(A)

Nectar removal plot

Control plot

(B)

Nectar removal plot

Control plot

biological knowledge as possible. Try to look at the world from the organism’s perspective to think about how far to separate samples. Pilot studies, if feasible, also can suggest appropriate spacing to ensure independence. So why not just maximize the distance or time between samples? First, as we described earlier, it becomes more expensive to collect data as the distance between samples increases. Second, moving the samples very far apart can introduce new sources of variation because of differences (heterogeneity) within or among habitats. We want our replicates close enough together to ensure we are sampling relatively hom*ogenous or consistent conditions, but far enough apart to ensure that the responses we measure are independent of one another. In spite of its central importance, the independence problem is almost never discussed explicitly in scientific papers. In the Methods section of a paper, you are likely to read a sentence such as, “We measured 100 randomly selected seedlings growing in full sunlight. Each measured seedling was at least 50 cm from its nearest neighbor.” What the authors mean is, “We don’t know how far apart the observations would have to have been in order to ensure independence. However, 50 cm seemed like a fair distance for the tiny seedlings we studied. If we had chosen distances greater than 50 cm, we could not have collected all of our data in full sunlight, and some of the seedlings would have been collected in the shade, which obviously would have influenced our results.”

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Avoiding Confounding Factors

Avoiding Confounding Factors When factors are confounded with one another, their effects cannot be easily disentangled. Let’s return to the hummingbird example. Suppose we prudently separated the control and nectar removal plots, but inadvertently placed the removal plot on a sunny hillside and the control plot in a cool valley (Figure 6.6). Hummingbirds forage less frequently in the removal plot (7 visits/hour), and the two plots are now far enough apart that there is no problem of independence. However, hummingbirds naturally tend to avoid foraging in the cool valley, so the foraging rate also is low in these plots (6 visits/hour). Because the treatments are confounded with temperature differences, we cannot tease apart the effects of foraging preferences from those of thermal preferences. In this case, the two forces largely cancel one another, leading to comparable foraging rates in the two plots, although for very different reasons. This example may seem a bit contrived. Knowing the thermal preferences of hummingbirds, we would not have set up such an experiment. The problem is that there are likely to be unmeasured or unknown variables—even in an apparently hom*ogenous environment—that can have equally strong effects on our experiment. And, if we are conducting a natural experiment, we are stuck with whatever confounding factors are present in the environment. In an observational study of hummingbird foraging, we may not be able to find plots that differ only in their levels of nectar rewards but do not also differ in temperature and other factors known to affect foraging behavior.

Nectar removal plot

Control plot

Warmer

Cooler

Figure 6.6 A confounded experimental design. As in Figure 6.5, the study establishes control and experimental nectar removal plots to evaluate the responses of foraging hummingbirds. In this design, although the plots are far enough apart to ensure independence, they have been placed at different points along a thermal gradient. Consequently, the treatment effects are confounded with differences in the thermal environment. The net result is that the experiment compares data from a warm nectar removal plot with data from a cool control plot.

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Replication and Randomization The dual threats of confounding factors and non-independence would seem to threaten all of our statistical conclusions and render even our experimental studies suspect. Incorporating replication and randomization into experimental designs can largely offset the problems introduced by confounding factors and non-independence. By replication, we mean the establishment of multiple plots or observations within the same treatment or comparison group. By randomization, we mean the random assignment of treatments or selection of samples.9 Let’s return one more time to the hummingbird example. If we follow the principles of randomization and replication, we will set up many replicate control and removal plots (ideally, a minimum of 10 of each). The location of each of these plots in the study area will be random, and the assignment of the treatment (control or removal) to each plot also will be random (Figure 6.7).10 How will randomization and replication reduce the problem of confounding factors? Both the warm hillside, the cool valley, and several intermediate sites each will have multiple plots from both control and nectar removal treatments. Thus, the temperature factor is no longer confounded with the treatment, as all treatments occur within each level of temperature. As an additional benefit, this design will also allow you to test the effects of temperature as a covariate on hummingbird foraging behavior, independent of the levels of nectar (see Chapters 7 and 10). It is true that hummingbird visits will still be more frequent on the warm hillside than in the cool valley, but that will be true for replicates of both the control and nectar removal. The temperature will add more variation to the data, but it will not bias the results because the warm and cool plots will

Many samples that are claimed to be random are really haphazard. Truly random sampling means using a random number generator (such as the flip of a fair coin, the roll of a fair die, or the use of a reliable computer algorithm for producing random numbers) to decide which replicates to use. In contrast, with haphazard sampling, an ecologist follows a set of general criteria [e.g., mature trees have a diameter of more than 3 cm at breast height (dbh = 1.3 m)] and selects sites or organisms that are spaced hom*ogenously or conveniently within a sample area. Haphazard sampling is often necessary at some level because random sampling is not efficient for many kinds of organisms, especially if their distribution is spatially patchy. However, once a set of organisms or sites is identified, randomization should be used to sample or to assign replicates to different treatment groups. 10 Randomization takes some time, and you should do as much of it as possible in advance, before you get into the field. It is easy to generate random numbers and simulate random sampling with computer spreadsheets. But it is often the case that you will need to generate random numbers in the field. Coins and dice (especially 10-sided

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Replication and Randomization

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Figure 6.7 A properly replicated and randomized experimental design. The study establishes plots as in Figures 6.6. Each square represents a replicate control plot (black dots) or nectar removal plot (gray dots). The plots are separated by enough distance to ensure independence, but their location within the temperature gradient has been randomized. There are 10 replicates for each of the two treatments. The spatial scale of the drawing is larger than in Figure 6.6. Warmer

Cooler

be distributed approximately equally between the control and removal treatments. Of course, if we knew ahead of time that temperature was an important determinant of foraging behavior, we might not have used this design for the experiment. Randomization minimizes the confounding of treatments with unknown or unmeasured variables in the study area. It is less obvious how randomization and replication reduce the problem of non-independence among samples. After all, if the plots are too close together, the foraging visits will not be independent, regardless of the amount of replication or randomization. Whenever possible, we should use common sense and

gaming dice) are useful for this purpose. A clever trick is to use a set of coins as a binary random number generator. For example, suppose you have to assign each of your replicates to one of 8 different treatments, and you want to do so randomly. Toss 3 coins, and convert the pattern of heads and tails to a binary number (i.e., a number in base 2). Thus, the first coin indicates the 1s, the second coin indicates the 2s, the third coin indicates the 4s, and so on. Tossing 3 coins will give you a random integer between 0 and 7. If your three tosses are heads, tails, heads (HTH), you have a 1 in the one’s place, a 0 in the two’s place, and a 1 in the four’s place. The number is 1 + 0 + 4 = 5. A toss of (THT) is 0 + 2 + 0 = 2. Three tails gives you a 0 (0 + 0 + 0) and three heads give you a 7 (1 + 2 + 4). Tossing 4 coins will give you 16 integers, and 5 coins will give you 32. An even easier method is to take a digital stopwatch into the field. Let the watch run for a few seconds and then stop it without looking at it. The final digit that measures time in 1/100th of a second can be used as a random uniform digit from 0 to 9. A statistical analysis of 100 such random digits passed all of the standard diagnostic tests for randomness and uniformity (B. Inouye, personal communication).

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knowledge of biology to separate plots or samples by some minimum distance or sampling interval to avoid dependence. However if we do not know all of the forces that can cause dependence, a random placement of plots beyond some minimum distance will ensure that the spacing of the plots is variable. Some plots will be relatively close, and some will be relatively far apart. Therefore, the effect of the dependence will be strong in some pairs of plots, weak in others, and nonexistent in still others. Such variable effects may cancel one another and can reduce the chances that results are consistently biased by non-independence. Finally, note that randomization and replication only are effective when they are used together. If we do not replicate, but simply assign randomly the control and treatment plots to the hillside or the valley, the design is still confounded (see Figure 6.6). Similarly, if we replicate the design, but assign all 10 of the controls to the valley and all 10 of the removals to the hillside, the design is also confounded (Figure 6.8). It is only when we use multiple plots and assign the treatments randomly that the confounding effect of temperature is removed from the design (see Figure 6.7). Indeed, it is fair to say that any unreplicated design is always going to be confounded with one or more environmental factors.11 Although the concept of randomization is straightforward, it must be applied at several stages in the design. First, randomization applies only to a well-defined, initially non-random sample space. The sample space doesn’t simply mean the physical area from which replicates are sampled (although this is an important aspect of the sample space). Rather, the sample space refers to a set of elements that have experienced similar, though not identical, conditions. Examples of a sample space might include individual cutthroat trout that are reproductively mature, lightfall gaps created by fires, old-fields abandoned 10–20

11 Although confounding is easy to recognize in a field experiment of this sort, it may not be apparent that the same problem exists in laboratory and greenhouse experiments. If we rear insect larvae at high and low temperatures in two environmental chambers, this is a confounded design because all of the high temperature larvae are in one chamber and all of the low temperature larvae are in the other. If environmental factors other than temperature also differ between the chambers, their effects are confounded with temperature. The correct solution would be to rear each larva in its own separate chamber, thereby ensuring that each replicate is truly independent and that temperature is not confounded with other factors. But this sort of design simply is too expensive and wasteful of space ever to be used. Perhaps the argument can be made that environmental chambers and greenhouses really do differ only in temperature and no other factors, but that is only an assumption that should be tested explicitly. In many cases, the environment in environmental chambers is surprisingly heterogeneous, both within and between chambers. Potvin (2001) discusses how this variability can be measured and then used to design better laboratory experiments.

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Replication and Randomization

Figure 6.8 A replicated, but confounded, design. As in Figures 6.5, 6.6, and 6.7, the study establishes control and experimental nectar removal plots to evaluate the responses of foraging hummingbirds. Each square represents a replicate control plot (black dots) or nectar removal plot (gray dots). If treatments are replicated but not assigned randomly, the design still confounds treatments with underlying environmental gradients. Replication combined with randomization and sufficient spacing of replicates (see Figure 6.7) is the only safeguard against non-independence (see Figure 6.5) and confounding (see Figures 6.6 and 6.8).

years ago, or large bleached coral heads at 5–10 meters depth. Once this sample space has been defined clearly, sites, individuals, or replicates that meet the criteria should be chosen at random. As we noted in Chapter 1, the spatial and temporal boundaries of the study will dictate not only the sampling effort involved, but also the domain of inference for the conclusions of the study. Once sites or samples are randomly selected, treatments should be assigned to them randomly, which ensures that different treatments are not clumped in space or confounded with environmental variables.12 Samples should also be collected and treatments applied in a random sequence. That way, if environmental conditions change during the experiment, the results will not be

If the sample size is too small, even a random assignment can lead to spatial clumping of treatments. One solution would be to set out the treatments in a repeated order (…123123…), which ensures that there is no clumping. However, if there is any nonindependence among treatments, this design may exaggerate its effects, because Treatment 2 will always occur spatially between Treatments 1 and 3. A better solution would be to repeat the randomization and then statistically test the layout to ensure there is no clumping. See Hurlbert (1984) for a thorough discussion of the numerous hazards that can arise by failing to properly replicate and randomize ecological experiments.

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confounded. For example, if you census all of your control plots first, and your field work is interrupted by a fierce thunderstorm, any impacts of the storm will be confounded with your manipulations because all of the treatment plots will be censused after the storm. These same provisos hold for non-experimental studies in which different plots or sites have to be censused. The caveat is that strictly random censusing in this way may be too inefficient because you will usually not be visiting neighboring sites in consecutive order. You may have to compromise between strict randomization and constraints imposed by sampling efficiency. All methods of statistical analysis—whether they are parametric, Monte Carlo, or Bayesian (see Chapter 5)—rest on the assumption of random sampling at an appropriate spatial or temporal scale. You should get in the habit of using randomization whenever possible in your work.

Designing Effective Field Experiments and Sampling Studies Here are some questions to ask when designing field experiments and sampling studies. Although some of these questions appear to be specific to manipulative experiments, they are also relevant to certain natural experiments, where “controls” might consist of plots lacking a particular species or set of abiotic conditions. Are the Plots or Enclosures Large Enough to Ensure Realistic Results?

Field experiments that seek to control animal density must necessarily constrain the movement of animals. If the enclosures are too small, the movement, foraging, and mating behaviors of the animals may be so unrealistic that the results obtained will be uninterpretable or meaningless (MacNally 2000a). Try to use the largest plots or cages that are feasible and that are appropriate for the organism you are studying. The same considerations apply to sampling studies: the plots need to be large enough and sampled at an appropriate spatial scale to answer your question. What Is the Grain and Extent of the Study?

Although much importance has been placed on the spatial scale of an experiment or a sampling study, there are actually two components of spatial scale that need to be addressed: grain and extent. Grain is the size of the smallest unit of study, which will usually be the size of an individual replicate or plot. Extent is the total area encompassed by all of the sampling units in the study. Grain and extent can be either large or small (Figure 6.9). There is no single combination of grain and extent that is necessarily correct. However, ecological studies with

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Designing Effective Field Experiments and Sampling Studies

Spatial extent

Small

Large

Small

Large Spatial grain

Figure 6.9 Spatial grain and spatial extent in ecological studies. Each square represents a single plot. Spatial grain measures the size of the sampling units, represented by small or large squares. Spatial extent measures the area encompassing all of the replicates of the study, represented by closely grouped or widely spaced squares.

both a small grain and a small extent, such as pitfall catches of beetles in a single forest plot, may sometimes be too limited in scope to allow for broad conclusions. On the other hand, studies with a large grain but a small extent, such as whole-lake manipulations in a single valley, may be very informative. Our own preference is for studies with a small grain, but a medium or large extent, such as ant and plant censuses in small plots (5 m × 5 m) across New England (Gotelli and Ellison 2002a,b) or eastern North America (Gotelli and Arnett 2000), or on small mangrove islands in the Caribbean (Farnsworth and Ellison 1996a). The small grain allows for experimental manipulations and observations taken at scales that are relevant to the organism, but the large extent expands the domain of inference for the results. In determining grain and extent, you should consider both the question you are trying to ask and the constraints on your sampling. Does the Range of Treatments or Census Categories Bracket or Span the Range of Possible Environmental Conditions?

Many field experiments describe their manipulations as “bracketing or spanning the range of conditions encountered in the field.” However, if you are trying to model climate change or altered environments, it may be necessary to also include conditions that are outside the range of those normally encountered in the field.

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Have Appropriate Controls Been Established to Ensure that Results Reflect Variation Only in the Factor of Interest?

It is rare that a manipulation will change one and only one factor at a time. For example, if you surround plants with a cage to exclude herbivores, you have also altered the shading and moisture regime. If you simply compare these plants to unmanipulated controls, the herbivore effects are confounded with the differences in shading and moisture. The most common mistake in experimental designs is to establish a set of unmanipulated plots and then treat those as a control. Usually, an additional set of control plots that contain some minimal alteration will be necessary to properly control for the manipulations. In the example described above, an open-sided cage roof will allow herbivores access to plants, but will still include the shading effects of the cage. With this simple design of three treatments (Unmanipulated, Cage control, Herbivore exclusion), you can make the following contrasts: 1. Unmanipulated versus Cage control. This comparison reveals the extent to which shading and physical changes due to the cage per se are affecting plant growth and responses. 2. Cage control versus Herbivore exclusion. This comparison reveals the extent to which herbivory alters plant growth. Both the Control and Herbivore exclusion plots experience the shading effects of the cage, so any difference between them can be attributed to the effect of herbivores. 3. Unmanipulated versus Herbivore exclusion. This comparison measures the combined effect of both the herbivores and the shading on plant growth. Because the experiment is designed to measure only the herbivore effect, this particular comparison confounds treatment and caging effects. In Chapter 10, we will explain how you use can use contrasts after analysis of variance to quantify these comparisons. Have All Replicates Been Manipulated in the Same Way Except for the Intended Treatment Application?

Again, appropriate controls usually require more than lack of manipulation. If you have to push back plants to apply treatments, you should push back plants in the control plots as well (Salisbury 1963; Jaffe 1980). In a reciprocal transplant experiment with insect larvae, live animals may be sent via overnight courier to distant sites and established in new field populations. The appropriate control is a set of animals that are re-established in the populations from which they were collected. These animals will also have to receive the “UPS treatment” and be sent through the mail system to ensure they receive the same stress as the ani-

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Summary

mals that were transplanted to distant sites. If you are not careful to ensure that all organisms are treated identically in your experiments, your treatments will be confounded with differences in handling effects (Cahill et al. 2000). Have Appropriate Covariates Been Measured in Each Replicate?

Covariates are continuous variables (see Chapter 7) that potentially affect the response variable, but are not necessarily controlled or manipulated by the investigator. Examples include variation among plots in temperature, shade, pH, or herbivore density. Different statistical methods, such as analysis of covariance (see Chapter 10), can be used to quantify the effect of covariates. However, you should avoid the temptation to measure every conceivable covariate in a plot just because you have the instrumentation (and the time) to do so. You will quickly end up with a dataset in which you have more variables measured than you have replicates, which causes additional problems in the analysis (Burnham and Anderson 2010). It is better to choose ahead of time the most biologically relevant covariates, measure only those covariates, and use sufficient replication. Remember also that the measurement of covariates is useful, but it is not a substitute for proper randomization and replication.

Summary The sound design of an ecological experiment first requires a clear statement of the question being asked. Both manipulative and observational experiments can answer ecological questions, and each type of experiment has its own strengths and weaknesses. Investigators should consider the appropriateness of using a press versus a pulse experiment, and whether the replication will be in space (snapshot experiment), time (trajectory experiment), or both. Non-independence and confounding factors can compromise the statistical analysis of data from both manipulative and observational studies. Randomization, replication, and knowledge of the ecology and natural history of the organisms are the best safeguards against non-independence and confounding factors. Whenever possible, try to use at least 10 observations per treatment group. Field experiments usually require carefully designed controls to account for handling effects and other unintended alterations. Measurement of appropriate environmental covariates can be used to account for uncontrolled variation, although it is not a substitute for randomization and replication.

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CHAPTER 7

A Bestiary of Experimental and Sampling Designs

In an experimental study, we have to decide on a set of biologically realistic manipulations that include appropriate controls. In an observational study, we have to decide which variables to measure that will best answer the question we have asked. These decisions are very important, and were the subject of Chapter 6. In this chapter we discuss specific designs for experimental and sampling studies in ecology and environmental science. The design of an experiment or observational study refers to how the replicates are physically arranged in space, and how those replicates are sampled through time. The design of the experiment is intimately linked to the details of replication, randomization, and independence (see Chapter 6). Certain kinds of designs have proven very powerful for the interpretation and analysis of field data. Other designs are more difficult to analyze and interpret. However, you cannot draw blood from a stone, and even the most sophisticated statistical analysis cannot rescue a poor design. We first present a simple framework for classifying designs according to the types of independent and dependent variables. Next, we describe a small number of useful designs in each category. We discuss each design and the kinds of questions it can be used to address, illustrate it with a simple dataset, and describe the advantages and disadvantages of the design. The details of how to analyze data from these designs are postponed until Chapters 9–12. The literature on experimental and sampling designs is vast (e.g., Cochran and Cox 1957; Winer 1991; Underwood 1997; Quinn and Keough 2002), and we present only a selective coverage in this chapter. We restrict ourselves to those designs that are practical and useful for ecologists and environmental scientists, and that have proven to be most successful in field studies.

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Categorical versus Continuous Variables We first distinguish between categorical variables and continuous variables. Categorical variables are classified into one of two or more unique categories. Ecological examples include sex (male, female), trophic status (producer, herbivore, carnivore), and habitat type (shade, sun). Continuous variables are measured on a continuous numerical scale; they can take on a range of real number or integer values. Examples include measurements of individual size, species richness, habitat coverage, and population density. Many statistics texts make a further distinction between purely categorical variables, in which the categories are not ordered, and ranked (or ordinal) variables, in which the categories are ordered based on a numerical scale. An example of an ordinal variable would be a numeric score (0, 1, 2, 3, or 4) assigned to the amount of sunlight reaching the forest floor: 0 for 0–5% light; 1 for 6–25% light; 2 for 26–50% light; 3 for 51–75% light; and 4 for 76–100% light. In many cases, methods used for analyzing continuous data also can be applied to ordinal data. In a few cases, however, ordinal data are better analyzed with Monte Carlo methods, which were discussed in Chapter 5. In this book, we use the term categorical variable to refer to both ordered and unordered categorical variables. The distinction between categorical and continuous variables is not always clear-cut; in many cases, the designation depends simply on how the investigator chooses to measure the variable. For example, a categorical habitat variable such as sun/shade could be measured on a continuous scale by using a light meter and recording light intensity in different places. Conversely, a continuous variable such as salinity could be classified into three levels (low, medium, and high) and treated as a categorical variable. Recognizing the kind of variable you are measuring is important because different designs are based on categorical and continuous variables. In Chapter 2, we distinguished two kinds of random variables: discrete and continuous. What’s the difference between discrete and continuous random variables on the one hand, and categorical and continuous variables on the other? Discrete and continuous random variables are mathematical functions for generating values associated with probability distributions. In contrast, categorical and continuous variables describe the kinds of data that we actually measure in the field or laboratory. Continuous variables usually can be modeled as continuous random variables, whereas both categorical and ordinal variables usually can be modeled as discrete random variables. For example, the categorical variable sex can be modeled as a binomial random variable; the numerical variable height can be modeled as normal random variable; and the ordinal variable light reaching the forest floor can be modeled as a binomial, Poisson, or uniform random variable.

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Four Classes of Experimental Design

Dependent and Independent Variables After identifying the types of variables with which you are working, the next step is to designate dependent and independent variables. The assignment of dependent and independent variables implies an hypothesis of cause and effect that you are trying to test. The dependent variable is the response variable that you are measuring and for which you are trying to determine a cause or causes. In a scatterplot of two variables, the dependent or response variable is called the Y variable, and it usually is plotted on the ordinate (vertical or y-axis). The independent variable is the predictor variable that you hypothesize is responsible for the variation in the response variable. In the same scatterplot of two variables, the independent or predictor variable is called the X variable, and it usually is plotted on the abscissa (horizontal or x-axis).1 In an experimental study, you typically manipulate or directly control the levels of the independent variable and measure the response in the dependent variable. In an observational study, you depend on natural variation in the independent variable from one replicate to the next. In both natural and experimental studies, you don’t know ahead of time the strength of the predictor variable. In fact, you are often testing the statistical null hypothesis that variation in the response variable is unrelated to variation in the predictor variable, and is no greater than that expected by chance or sampling error. The alternative hypothesis is that chance cannot entirely account for this variation, and that at least some of the variation can be attributed to the predictor variable. You also may be interested in estimating the size of the effect of the predictor or causal variable on the response variable.

Four Classes of Experimental Design By combining variable types—categorical versus continuous, dependent versus independent—we obtain four different design classes (Table 7.1). When independent variables are continuous, the classes are either regression (continuous dependent variables) or logistic regression (categorical dependent variables). When independent variables are categorical, the classes are either ANOVA (continuous dependent variable) or tabular (categorical dependent variable). Not all designs fit nicely into these four categories. The analysis of covariance

Of course, merely plotting a variable on the x-axis is does not guarantee that it is actually the predictor variable. Particularly in natural experiments, the direction of cause and effect is not always clear, even though the measured variables may be highly correlated (see Chapter 6).

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TABLE 7.1 Four classes of experimental and sampling designs Dependent variable Continuous Categorical

Independent variable Continuous

Categorical

Regression Logistic regression

ANOVA Tabular

Different kinds of designs are used depending on whether the independent and dependent variables are continuous or categorical. When both the dependent and the independent variables are continuous, a regression design is used. If the dependent variable is categorical and the independent variable is continuous, a logistic regression design is used. The analysis of regression designs is covered in Chapter 9. If the independent variable is categorical and the dependent variable is continuous, an analysis of variance (ANOVA) design is used. The analysis of ANOVA designs is described in Chapter 10. Finally, if both the dependent and independent variables are categorical, a tabular design is used. Analysis of tabular data is described in Chapter 11.

(ANCOVA) is used when there are two independent variables, one of which is categorical and one of which is continuous (the covariate). ANCOVA is discussed in Chapter 10. Table 7.1 categorizes univariate data, in which there is a single dependent variable. If, instead, we have a vector of correlated dependent variables, we rely on a multivariate analysis of variance (MANOVA) or other multivariate methods that are described in Chapter 12. Regression Designs

When independent variables are measured on continuous numerical scales (see Figure 6.1 for an example), the sampling layout is a regression design. If the dependent variable is also measured on a continuous scale, we use linear or nonlinear regression models to analyze the data. If the dependent variable is measured on an ordinal scale (an ordered response), we use logistic regression to analyze the data. These three types of regression models are discussed in detail in Chapter 9. SINGLE-FACTOR REGRESSION

A regression design is simple and intuitive. Collect data on a set of independent replicates. For each replicate, measure both the predictor and the response variables. In an observational study, neither of the two variables is manipulated, and your sampling is dictated by the levels of natural variation in the independent variable. For example, suppose your hypothesis is that the density of desert rodents is controlled by the availability of seeds (Brown and Leiberman 1973). You could sample 20 independent plots, each

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Four Classes of Experimental Design

chosen to represent a different abundance level of seeds. In each plot, you measure the density of seeds and the density of desert rodents (Figure 7.1). The data are organized in a spreadsheet in which each row is a different plot, and each column is a different response or predictor variable. The entries in each row represent the measurements taken in a single plot. In an experimental study, the levels of the predictor variable are controlled and manipulated directly, and you measure the response variable. Because your hypothesis is that seed density is responsible for desert rodent density (and not the other way around), you would manipulate seed density in an experimental study, either adding or removing seeds to alter their availability to rodents. In both the experimental study and the observational study, your assumption is that the predictor variable is a causal variable: changes in the value of the predictor (seed density) would cause a change in the value of the response (rodent density). This is very different from a study in which you would examine the correlation (statistical covariation) between the two variables. Correlation does not specify a cause-and-effect relationship between the two variables.2 In addition to the usual caveats about adequate replication and independence of the data (see Chapter 6), two principles should be followed in designing a regression study: 1. Ensure that the range of values sampled for the predictor variable is large enough to capture the full range of responses by the response variable. If the predictor variable is sampled from too limited a range, there may appear to be a weak or nonexistent statistical relationship between the predictor

The sampling scheme needs to reflect the goals of the study. If the study is designed simply to document the relationship between seeds and rodent density, then a series of random plots can be selected, and correlation is used to explore the relationship between the two variables. However, if the hypothesis is that seed density is responsible for rodent density, then a series of plots that encompass a uniform range of seed densities should be sampled, and regression is used to explore the functional dependence of rodent abundance on seed density. Ideally, the sampled plots should differ from one another only in the density of seeds present. Another important distinction is that a true regression analysis assumes that the value of the independent variable is known exactly and is not subject to measurement error. Finally, standard linear regression (also referred to as Model I regression) minimizes residual deviations in the vertical (y) direction only, whereas correlation minimizes the perpendicular (x and y) distance of each point from the regression line (also referred to as Model II regression). The distinction between correlation and regression is subtle, and is often confusing because some statistics (such as the correlation coefficient) are identical for both kinds of analyses. See Chapter 9 for more details.

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Plot 2 Rodent density: 1.1 Seed density: 1,500 Vegetation cover: 2

Plot number 1 2 . . 20

Seeds/m2 12,000 1,500 . . 11,500

Plot 1 Rodent density: 5.0 Seed density: 12,000 Vegetation cover: 11

Vegetation cover (%) 11 2 . . 52

Rodents/m2 5.0 1.1 . . 3.7

Figure 7.1 Spatial arrangement of replicates for a regression study. Each square represents a different 25-m2 plot. Plots were sampled to ensure a uniform coverage of seed density (see Figures 7.2 and 7.3). Within each plot, the investigator measures rodent density (the response variable), and seed density and vegetation cover (the two predictor variables). The data are organized in a spreadsheet in which each row is a plot, and the columns are the measured variables within the plot.

and response variables even though they are related (Figure 7.2). A limited sampling range makes the study susceptible to a Type II statistical error (failure to reject a false null hypothesis; see Chapter 4). 2. Ensure that the distribution of predictor values is approximately uniform within the sampled range. Beware of datasets in which one or two of the values of the predictor variable are very different in size from the others. These influential points can dominate the slope of the regression and

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(A)

Figure 7.2 Inadequate sampling over a narrow

Y variable

Four Classes of Experimental Design

range (within the dashed lines) of the X variable can create a spuriously non-significant regression slope, even though X and Y are strongly correlated with one another. Each point represents a single replicate for which a value has been measured for both the X and the Y variables. Blue circles represent possible data that were not collected for the analysis. Black circles represent the sample of replicates that were measured. (A) The full range of data. The solid line indicates the true linear relationship between the variables. (B) The regression line is fitted to the sample data. Because the X variable was sampled over a narrow range of values, there is limited variation in the resulting Y variable, and the slope of the fitted regression appears to be close to zero. Sampling over the entire range of the X variable will prevent this type of error.

Y variable

(B)

X variable

generate a significant relationship where one does not really exist (Figure 7.3; see Chapter 8 for further discussion of such outliers). Sometimes influential data points can be corrected with a transformation of the predictor variable (see Chapter 8), but we re-emphasize that analysis cannot rescue a poor sampling design. MULTIPLE REGRESSION The extension to multiple regression is straightforward. Two or more continuous predictor variables are measured for each replicate, along with the single response variable. Returning to the desert rodent example, you suspect that, in addition to seed availability, rodent density is also controlled by vegetation structure—in plots with sparse vegetation, desert rodents are vul-

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Figure 7.3 Failure to sample uniformly the entire range of a variable can lead to spurious results. As in Figure 7.2, each black point represents a single recorded observation; the blue points represent unobserved X, Y pairs. (A) The solid line indicates the true linear relationship between the variables. This relationship would have been revealed if the X variable had been sampled uniformly. (B) The regression line is fitted to the sample data alone (i.e., just the black points). Because only a single datum with a large value of X was measured, this point has an inordinate influence on the fitted regression line. In this case, the fitted regression line inaccurately suggests a positive relationship between the two variables.

(A)

Y variable

170

Y variable

(B)

X variable

nerable to avian predators (Abramsky et al. 1997). In this case, you would take three measurements in each plot: rodent density, seed density, and vegetation cover. Rodent density is still the response variable, and seed density and vegetation cover are the two predictor variables (see Figure 7.1). Ideally, the different predictor variables should be independent of one another. As in simple regression designs, the different values of the predictor variables should be established evenly across the full range of possible values. This is straightforward in an experimental study, but rarely is achievable in an observational study. In an observational study, it is often the case that the predictor variables themselves will be correlated with each other. For example, plots with high vegetation density are likely to have high seed density. There may be few or no plots in which

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vegetation density is high and seed density is low (or vice versa). This collinearity makes it difficult to estimate accurately regression parameters3 and to tease apart how much variation in the response variable is actually associated with each of the predictor variables. As always, replication becomes important as we add more predictor variables to the analysis. Following the Rule of 10 (see Chapter 6), you should try to obtain at least 10 replicates for each predictor variable in your study. But in many studies, it is a lot easier to measure additional predictor variables than it is to obtain additional independent replicates. However, you should avoid the temptation to measure everything that you can just because it is possible. Try to select variables that are biologically important and relevant to the hypothesis or question you are asking. It is a mistake to think that a model selection algorithm, such as stepwise multiple regression, can identify reliably the “correct” set of predictor variables from a large dataset (Burnham and Anderson 2010). Moreover, large datasets often suffer from multicollinearity: many of the predictor variables are correlated with one another (Graham 2003). ANOVA Designs

If your predictor variables are categorical (ordered or unordered) and your response variables are continuous, your design is called an ANOVA (for analysis of variance). ANOVA also refers to the statistical analysis of these types of designs (see Chapter 10). ANOVA is rife with terminology. Treatments refer to the different categories of the predictor variables that are used. In an experimental study, the treatments represent the different manipulations that have been performed. In an observational study, the treatments represent the different groups that are being compared. The number of treatments in a study equals the number of categories being compared. Within each treatment, multiple observations will be made, and each of these observations is a replicate. In standard ANOVA designs, each replicate should be independent, both statistically and biologically, of the other replicates within and among treatments. Later in this

TERMINOLOGY

In fact, if one of the predictor variables can be described as a perfect linear function of the other one, it is not even algebraically possible to solve for the regression coefficients. Even when the problem is not this severe, correlations among predictor variables make it difficult to test and compare models. See MacNally (2000b) for a discussion of correlated variables and model-building in conservation biology.

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chapter, we will discuss certain ANOVA designs that relax the assumption of independence among replicates. We also distinguish between single-factor designs and multifactor designs. In a single-factor design, each of the treatments represents variation in a single predictor variable or factor. Each value of the factor that represents a particular treatment is called a treatment level. For example, a single-factor ANOVA design could be used to compare growth responses of plants raised at 4 different levels of nitrogen, or the growth responses of 5 different plant species to a single level of nitrogen. The treatment groups may be ordered (e.g., 4 nitrogen levels) or unordered (e.g., 5 plant species). In a multifactor design, the treatments cover two (or more) different factors, and each factor is applied in combination in different treatments. In a multifactor design, there are different levels of the treatment for each factor. As in the single-factor design, the treatments within each factor may be either ordered or unordered. For example, a two-factor ANOVA design would be necessary if you wanted to compare the responses of plants to 4 levels of nitrogen (Factor 1) and 4 levels of phosphorus (Factor 2). In this design, each of the 4 × 4 = 16 treatment levels represents a different combination of nitrogen level and phosphorus level. Each combination of nutrients is applied to all of the replicates within the treatment (Figure 7.4). Although we will return to this topic later, it is worth asking at this point what the advantage is of using a two-factor design. Why not just run two separate experiments? For example, you could test the effects of phosphorus in a oneway ANOVA design with 4 treatment levels, and you could test the effects of nitrogen in a separate one-way ANOVA design, also with 4 treatment levels. What is the advantage of using a two-way design with 16 phosphorus–nitrogen treatment combinations in a single experiment? One advantage of the two-way design is efficiency. It is likely to be more costeffective to run a single experiment—even one with 16 treatments—than to run two separate experiments with 4 treatments each. A more important advantage is that the two-way design allows you to test both for main effects (e.g., the effects of nitrogen and phosphorus on plant growth) and for interaction effects (e.g., interactions between nitrogen and phosphorus). The main effects are the additive effects of each level of one treatment averaged over all of the levels of the other treatment. For example, the additive effect of nitrogen would represent the response of plants at each nitrogen level, averaged over the responses to the phosphorus levels. Conversely, the additive effect of phosphorus would be measured as the response of plants at each phosphorus level, averaged over the responses to the different nitrogen levels.

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0.00 mg 10

Nitrogen treatment (one-way layout) 0.10 mg 0.50 mg 1.00 mg 10 10 10

Phosphorous treatment (one-way layout) 0.00 mg 0.05 mg 0.10 mg 0.25 mg 10 10 10 10

(Simultaneous N and P treatments in a two-way layout) Phosphorus treatment

0.00 mg 0.05 mg 0.10 mg 0.25 mg

0.00 mg 10 10 10 10

Nitrogen treatment 0.10 mg 0.50 mg 10 10 10 10 10 10 10 10

1.00 mg 10 10 10 10

Figure 7.4 Treatment combinations in single-factor designs (upper two panels) and in a two-factor design (lower panel). In all designs, the number in each cell indicates the number of independent replicate plots to be established. In the two single-factor designs (oneway layouts), the four treatment levels represent four different nitrogen or phosphorous concentrations (mg/L). The total sample size is 40 plots in each single-factor experiment. In the two-factor design, the 4 × 4 = 16 treatments represent different combinations of nitrogen and phosphorous concentrations that are applied simultaneously to a replicate plot. This fully crossed two-factor ANOVA design with 10 replicates per treatment combination would require a total sample size of 160 plots. See Figures 7.9 and 7.10 for other examples of a crossed two-factor design.

Interaction effects represent unique responses to particular treatment combinations that cannot be predicted simply from knowing the main effects. For example, the growth of plants in the high nitrogen–high phosphorus treatment might be synergistically greater than you would predict from knowing the simple additive effects of nitrogen and phosphorus at high levels. Interaction effects are frequently the most important reason for using a factorial design. Strong interactions are the driving force behind much ecological and evolutionary change, and often are more important than the main effects. Chapter 10 will discuss analytical methods and interaction terms in more detail. SINGLE-FACTOR ANOVA

The single-factor ANOVA is one of the simplest, but most powerful, experimental designs. After describing the basic one-way layout, we also explain the randomized block and nested ANOVA designs. Strictly speaking, the randomized block and nested ANOVA are two-factor designs, but the second factor (blocks, or subsamples) is included only to control for sampling variation and is not of primary interest.

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The one-way layout is used to compare means among two or more treatments or groups. For example, suppose you want to determine whether the recruitment of barnacles in the intertidal zone of a shoreline is affected by different kinds of rock substrates (Caffey 1982). You could start by obtaining a set of slate, granite, and concrete tiles. The tiles should be identical in size and shape, and differ only in material. Following the Rule of 10 (see Chapter 6), set out 10 replicates of each substrate type (N = 30 total). Each replicate is placed in the mid-intertidal zone at a set of spatial coordinates that were chosen with a random number generator (Figure 7.5). After setting up the experiment, you return 10 days later and count the number of new barnacle recruits inside a 10 cm × 10 cm square centered in the middle of each tile. The data are organized in a spreadsheet in which each row is a

Figure 7.5 Example of a oneway layout. This experiment is designed to test for the effect of substrate type on barnacle recruitment in the rocky intertidal (Caffey 1982). Each circle represents an independent rock substrate. There are 10 randomly placed replicates of each of three treatments, represented by the three shades of blue. The number of barnacle recruits is sampled from a 10-cm square in the center of each rock surface. The data are organized in a spreadsheet in which each row is an independent replicate. The columns indicate the ID number of each replicate (1–30), the treatment group (Cement, Slate, or Granite), the replicate number within each treatment (1–10), and the number of barnacle recruits (the response variable).

ID number: 1 Treatment: Granite Number of barnacle recruits: 12

ID number 1 2 3 4 5 6 7 8 9 . . 30

Treatment Granite Slate Cement Granite Slate Cement Granite Slate Cement . . Cement

ID number: 3 Treatment: Cement Number of barnacle recruits: 3 ID number: 8 Treatment: Slate Number of barnacle recruits: 11

Replicate 1 1 1 2 2 2 3 3 3 . . 10

Number of barnacle recruits 12 10 3 14 10 8 11 11 7 . . 8

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replicate. The first few columns contain identifying information associated with the replicate, and the last column of the spreadsheet gives the number of barnacles that recruited into the square. Although the details are different, this is the same layout used in the study of ant density described in Chapter 5: multiple, independent replicate observations are obtained for each treatment or sampling group. The one-way layout is one of the simplest but most powerful experimental designs, and it can readily accommodate studies in which the number of replicates per treatment is not identical (unequal sample sizes). The one-way layout allows you to test for differences among treatments, as well as to test more specific hypotheses about which particular treatment group means are different and which are similar (see “Comparing Means” in Chapter 10). The major disadvantage of the one-way layout is that it does not explicitly accommodate environmental heterogeneity. Complete randomization of the replicates within each treatment implies that they will sample the entire array of background conditions, all of which may affect the response variable. On the one hand, this is a good thing because it means that the results of the experiment can be generalized across all of these environments. On the other hand, if the environmental “noise” is much stronger than the “signal” of the treatment, the experiment will have low power; the analysis may not reveal treatment differences unless there are many replicates. Other designs, including the randomized block and the two-way layout, can be used to accommodate environmental variability. A second, more subtle, disadvantage of the one-way layout is that it organizes the treatment groups along a single factor. If the treatments represent distinctly different kinds of factors, then a two-way layout should be used to tease apart main effects and interaction terms. Interaction terms are especially important because the effect of one factor often depends on the levels of another. For example, the pattern of recruitment onto different substrates may depend on the levels of a second factor (such as predator density). RANDOMIZED BLOCK DESIGNS One effective way to incorporate environmental heterogeneity is to modify the one-way ANOVA and use a randomized block design. A block is a delineated area or time period within which the environmental conditions are relatively hom*ogeneous. Blocks may be placed randomly or systematically in the study area, but they should be arranged so that environmental conditions are more similar within blocks than between them. Once the blocks are established, replicates will still be assigned randomly to treatments, but there is a restriction on the randomization: a single replicate from each of the treatments is assigned to each block. Thus, in a simple ran-

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domized block design, each block contains exactly one replicate of all the treatments in the experiment. Within each block, the placement of the treatment replicates should be randomized. Figure 7.6 illustrates the barnacle experiment laid out as a randomized block design. Because there are 10 replicates, there are

ID number: 6 Block: 2 Treatment: Cement Number of barnacle recruits: 8

ID number: 2 Block: 1 Treatment: Slate Number of barnacle recruits: 10 ID number: 1 Block: 1 Treatment: Granite Number of barnacle recruits: 12

Figure 7.6 Example of a randomized block design. The 10 replicates of each of the three treatments are grouped in blocks— physical groupings of one replicate of each of the three treatments. Both the placement of blocks and placement of treatments within blocks are randomized. Data organization in the spreadsheet is identical to the one-way layout (Figure 7.5), but the replicate column is replaced by a column indicating the block with which each replicate is associated.

ID number 1 2 3 4 5 6 7 8 9 . . 30

Treatment Granite Slate Cement Granite Slate Cement Granite Slate Cement . . Cement

Block 1 1 1 2 2 2 3 3 3 . . 10

Number of barnacle recruits 12 10 3 14 10 8 11 11 7 . . 8

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10 blocks (fewer, if you replicate within each block), and each block will contain one replicate of each of the three treatments. The spreadsheet layout for these data is the same as for the one-way layout, except the replicate column is now replaced by a column indicating the block. Each block should be small enough to encompass a relatively hom*ogenous set of conditions. However, each block must also be large enough to accommodate a single replicate of each of the treatments. Moreover, there must be room within the block to allow sufficient spacing between replicates to ensure their independence (see Figure 6.5). The blocks themselves also have to be far enough apart from one another to ensure independence of replicates among blocks. If there are geographic gradients in environmental conditions, then each block should encompass a small interval of the gradient. For example, there are strong environmental gradients along a mountainside, so we might set up an experiment with three blocks, one each at high, medium, and low elevation (Figure 7.7A). But it would not be appropriate to create three blocks that run “across the grain” from high to low elevation (Figure 7.7B); each block encompasses conditions that are too heterogeneous. In other cases, the environmental variation may be patchy, and the blocks should be arranged to reflect that patchiness. For example, if an experiment is being conducted in a wetland complex, each semi-isolated fen could be treated as a block. Finally, if the spatial organization of environmental heterogeneity is unknown, the blocks can be arranged randomly within the study area.4 The randomized block design is an efficient and very flexible design that provides a simple control for environmental heterogeneity. It can be used to control for environmental gradients and patchy habitats. As we will see in Chapter 10, when environmental heterogeneity is present, the randomized block design is more efficient than a completely randomized one-way layout, which may require a great deal more replication to achieve the same statistical power.

4 The randomized block design allows you to set up your blocks to encompass environmental gradients in a single spatial dimension. But what if the variation occurs in two dimensions? For example, suppose there is a north-to-south moisture gradient in a field, but also an east-to-west gradient in predator density. In such cases, more complex randomized block designs can be used. For example, the Latin square is a block design in which the n treatments are placed in the field in an n × n square; each treatment appears exactly once in every row and once in every column of the layout. Sir Ronald Fisher (see Footnote 5 in Chapter 5) pioneered these kinds of designs for agricultural studies in which a single field is partitioned and treatments applied to the contiguous subplots. These designs have not been used often by ecologists because the restrictions on randomization and layout are difficult to achieve in field experiments.

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Figure 7.7 Valid and invalid blocking designs. (A) Three properly oriented blocks, each encompassing a single elevation on a mountainside or other environmental gradient. Environmental conditions are more similar within than among blocks. (B) These blocks are oriented improperly, going “across the grain” of the elevational gradient. Conditions are as heterogeneous within the blocks as between them, so no advantage is gained by blocking.

(A) Valid blocking

(B) Invalid blocking High elevation

Low elevation

The randomized block design is also useful when your replication is constrained by space or time. For example, suppose you are running a laboratory experiment on algal growth with 8 treatments and you want to complete 10 replicates per treatment. However, you have enough space in your laboratory to run only 12 replicates at a time. What can you do? You should run the experiment in blocks, in which you set up a single replicate of each of the 8 treatments. After the result is recorded, you set up the experiment again (including another set of randomizations for treatment establishment and placement) and continue until you have accumulated 10 blocks. This design controls for inevitable changes in environmental conditions that occur in your laboratory through time, but still allows for appropriate comparison of treatments. In other cases, the limitation may not be space, but organisms. For example, in a study of mating behavior of fish, you may have to wait until you have a certain number of sexually mature fish before you can set up and run a single block of the experiment. In both examples, the randomized block design is the best safeguard against variation in background conditions during the course of your experiment. Finally, the randomized block design can be adapted for a matched pairs layout. Each block consists of a group of individual organisms or plots that have been deliberately chosen to be most similar in background characteristics. Each replicate in the group receives one of the assigned treatments. For example, in a simple experimental study of the effects of abrasion on coral growth, a pair of coral heads of similar size would be considered a single block. One of the coral heads would be randomly assigned to the control group, and the other would be assigned to the abrasion group. Other matched pairs would be chosen in the same way and the treatments applied. Even though the individuals in each

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pair are not part of a spatial or a temporal block, they are probably going to be more similar than individuals in other such blocks because they have been matched on the basis of colony size or other characteristics. For this reason, the analysis will use a randomized block design. The matched pairs approach is a very effective method when the responses of the replicates potentially are very heterogeneous. Matching the individuals controls for that heterogeneity, making it easier to detect treatment effects. There are four disadvantages to the randomized block design. The first is that there is a statistical cost to running the experiment with blocks. If the sample size is small and the block effect is weak, the randomized block design is less powerful than a simple one-way layout (see Chapter 10). The second disadvantage is that if the blocks are too small you may introduce non-independence by physically crowding the treatments together. As we discussed in Chapter 6, randomizing the placement of the treatments within the block will help with this problem, but won’t eliminate it entirely. The third disadvantage of the randomized block design is that if any of the replicates are lost, the data from that block cannot be used unless the missing values can be estimated indirectly. The fourth—and most serious—disadvantage of the randomized block design is that it assumes there is no interaction between the blocks and the treatments. The blocking design accounts for additive differences in the response variable and assumes that the rank order of the responses to the treatment does not change from one block to the next. Returning to the barnacle example, the randomized block model assumes that if recruitment in one of the blocks is high, all of the observations in that block will have elevated recruitment. However, the treatment effects are assumed to be consistent from one block to the next, so that the rank order of barnacle recruitment among treatments (Granite > Slate > Cement) is the same, regardless of any differences in the overall recruitment levels among blocks. But suppose that in some blocks recruitment is highest on the cement substrate and in other blocks it is highest on the granite substrate. In this case, the randomized block design may fail to properly characterize the main treatment effects. For this reason, some authors (Mead 1988; Underwood 1997) have argued that the simple randomized block design should not be used unless there is replication within blocks. With replication, the design becomes a two-factor analysis of variance, which we discuss below. Replication within blocks will indeed tease apart main effects, block effects, and the interaction between blocks and treatments. Replication will also address the problem of missing or lost data from within a block. However, ecologists often do not have the luxury of replication within blocks, particularly when the blocking factor is not of primary interest. The simple randomized block

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design (without replication) will at least capture the additive component (often the most important component) of environmental variation that would otherwise be lumped with pure error in a simple one-way layout. A nested design refers to any design in which there is subsampling within each of the replicates. We will illustrate it with the barnacle example. Suppose that, instead of measuring recruitment for a replicate in a single 10 cm× 10 cm square, you decided to take three such measurements for each of the 30 tiles in the study (Figure 7.8). Although the number of replicates has

NESTED DESIGNS

ID number: 2 Treatment: Granite Replicate number: 1 Subsample: 2 Number of barnacle recruits: 10

ID number: 1 Treatment: Granite Replicate number: 1 Subsample: 1 Number of barnacle recruits: 12

Figure 7.8 Example of a nested design. The study is the same as that shown in Figures 7.5 and 7.6. The layout is identical to the one-way layout in Figure 7.5, but here three subsamples are taken for each independent replicate. In the spreadsheet, an additional column is added to indicate the subsample number, and the total number of observations is increased from 30 to 90.

ID number 1 2 3 4 5 6 7 8 9 . . 90

Treatment Granite Granite Granite Slate Slate Slate Cement Cement Cement . . Cement

ID number: 3 Treatment: Granite Replicate number: 1 Subsample: 3 Number of barnacle recruits: 11

Replicate Number of number Subsample barnacle recruits 1 1 12 1 2 10 1 11 3 2 14 1 2 2 10 2 7 3 3 5 1 3 6 2 3 3 10 . . . . . . 3 30 6

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not increased, the number of observations has increased from 30 to 90. In the spreadsheet for these data, each row now represents a different subsample, and the columns indicate from which replicate and from which treatment the subsample was taken. This is the first design in which we have included subsamples that are clearly not independent of one another. What is the rationale for such a sampling scheme? The main reason is to increase the precision with which we estimate the response for each replicate. Because of the Law of Large Numbers (see Chapter 3), the more subsamples we use, the more precisely we will estimate the mean for each replicate. The increase in precision should increase the power of the test. There are three advantages to using a nested design. The first advantage, as we noted, is that subsampling increases the precision of the estimate for each replicate in the design. Second, the nested design allows you to test two hypotheses: first, is there variation among treatments? And, second, is there variation among the replicates within a treatment? The first hypothesis is equivalent to a one-way design that uses the subsample averages as the observation for each replicate. The second hypothesis is equivalent to a one-way design that uses the subsamples to test for differences among replicates within treatments.5 Finally, the nested design can be extended to a hierarchical sampling design. For example, you could, in a single study, census subsamples nested within replicates, replicates nested within intertidal zones, intertidal zones nested within shores, shores nested within regions, and even regions nested within continents (Caffey 1985). The reason for carrying out this kind of sampling is that the variation in the data can be partitioned into components that represent each of the hierarchical levels of the study (see Chapter 10). For example, you might be able to show that 80% of the variation in the data occurs at the level of intertidal zones within shores, but only 2% can be attributed to variation among shores within a region. This would mean that barnacle density varies strongly from the high to the low intertidal, but doesn’t vary much from one shoreline to the next. Such statements are useful for assessing the relative importance of different mechanisms in producing pattern (Petraitis 1998; see also Figure 4.6). Nested designs potentially are dangerous in that they are often analyzed incorrectly. One of the most serious and common mistakes in ANOVA is for 5

You can think of this second hypothesis as a one-way design at a lower hierarchical level. For example, suppose you used the data only from the four replicates of the granite treatment. Consider each replicate as a different “treatment” and each subsample as a different “replicate” for that treatment. The design is now a one-way design that compares the replicates of the granite treatment.

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investigators to treat each subsample as an independent replicate and analyze the nested design as a one-way design (Hurlbert 1984). The non-independence of the subsamples artificially boosts the sample size (by threefold in our example, in which we took three subsamples from each tile) and badly inflates the probability of a Type I statistical error (i.e., falsely rejecting a true null hypothesis). A second, less serious problem, is that the nested design can be difficult or even impossible to analyze properly if the sample sizes are not equal in each group. Even with equal numbers of samples and subsamples, nested sampling in more complex layouts, such as the two-way layout or the split-plot design, can be tricky to analyze; the simple default settings for statistical software usually are not appropriate. But the most serious disadvantage of the nested design is that it often represents a case of misplaced sampling effort. As we will see in Chapter 10, the power of ANOVA designs depends much more on the number of independent replicates than on the precision with which each replicate is measured. It is a much better strategy to invest your sampling effort in obtaining more independent replicates than subsampling within each replicate. By carefully specifying your sampling protocol (e.g., “only undamaged fruits from uncrowded plants growing in full shade”), you may be able to increase the precision of your estimates more effectively than by repeated subsampling. That being said, you should certainly go ahead and subsample if it is quick and cheap to do so. However, our advice is that you then average (or pool) those subsamples so that you have a single observation for each replicate and then treat the experiment as a one-way design. As long as the numbers aren’t too unbalanced, averaging also can alleviate problems of unequal sample size among subsamples and improve the fit of the errors to a normal distribution. It is possible, however, that after averaging among subsamples within replicates you no longer have sufficient replicates for a full analysis. In that case, you need a design with more replicates that are truly independent. Subsampling is no solution to the problem of inadequate replication! MULTIPLE-FACTOR DESIGNS: TWO-WAY LAYOUT

Multifactor designs extend the principles of the one-way layout to two or more treatment factors. Issues of randomization, layout, and sampling are identical to those discussed for the one-way, randomized block, and nested designs. Indeed, the only real difference in the design is in the assignment of the treatments to two or more factors instead of to a single factor. As before, the factors can represent either ordered or unordered treatments. Returning again to the barnacle example, suppose that, in addition to substrate effects, you wanted to test the effects of predatory snails on barnacle recruitment. You could set up a second one-way experiment in which you established

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four treatments: unmanipulated, cage control,6 predator exclusion, and predator inclusion. Instead of running two separate experiments, however, you decide to examine both factors in a single experiment. Not only is this a more efficient use of your field time, but also the effect of predators on barnacle recruitment might differ depending on the substrate type. Therefore, you establish treatments in which you simultaneously apply a different substrate and a different predation treatment. This is an example of a factorial design in which two or more factors are tested simultaneously in one experiment. The key element of a proper factorial design is that the treatments are fully crossed or orthogonal: every treatment level of the first factor (substrate) must be represented with every treatment level of the second factor (predation; Figure 7.9). Thus, the two-factor experiment has 3× 4 = 12 distinct treatment combinations, as opposed to only 3 treatments for the single-factor substrate experiment or 4 treatments for the single-factor predation experiment. Notice that each of these single-factor experiments would be restricted to only one of the treatment combinations of the other factor. In other words, the substrate experiment that we described above was conducted with the unmanipulated predation treatment, and the predation treatment would be conducted on only a single substrate type. Once we have determined the treatment combinations, the physical set up of the experiment would be the same as for a oneway layout with 12 treatment combinations (Figure 7.10). In the two-factor experiment, it is critical that all of the crossed treatment combinations be represented in the design. If some of the treatment combinations are missing, we end up with a confounded design. As an extreme example, suppose we set up only the granite substrate–predator exclusion treatment and the slate substrate–predator inclusion treatment. Now the predator effect is confounded with the substrate effect. Whether the results are statistically significant or not, we cannot tease apart whether the pattern is due to the effect of the predator, the effect of the substrate, or the interaction between them. This example highlights an important difference between manipulative experiments and observational studies. In the observational study, we would gather data on variation in predator and prey abundance from a range of samples. 6

Cage and cage control

In a cage control, investigators attempt to mimic the physical conditions generated by the cage, but still allow organisms to move freely in and out of the plot. For example, a cage control might consist of a mesh roof (placed over a plot) that allows predatory snails to enter from the sides. In an exclusion treatment, all predators are removed from a mesh cage, and in the inclusion treatment, predators are placed inside each mesh cage. The accompanying figure illustrates a cage (upper panel) and cage control (lower panel) in a fish exclusion experiment in a Venezuelan stream (Flecker 1996).

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Substrate treatment (one-way layout) Granite Slate Cement 10 10 10

Unmanipulated 10

Predator treatment (one-way layout) Control Predator exclusion Predator inclusion 10 10 10

(Simultaneous predator and substrate treatments in a two-way layout) Unmanipulated Predator treatment

Control Predator exclusion Predator inclusion

Granite 10

Substrate treatment Slate 10

Cement 10

Figure 7.9 Treatment combinations in two single-factor designs and in a fully crossed two-factor design. This experiment is designed to test for the effect of substrate type (Granite, Slate, or Cement) and predation (Unmanipulated, Control, Predator exclusion, Predator inclusion) on barnacle recruitment in the rocky intertidal. The number 10 indicates the total number of replicates in each treatment. The three shaded colors of the circles represent the three substrate treatments, and the patterns of the squares represent the four predation treatments. The two upper panels illustrate two one-way designs, in which only one of the two factors is systematically varied. In the two-factor design (lower panel), the 4 × 3 = 12 treatments represent different combinations of substrate and predation. The symbol in each cell indicates the combination of predation and substrate treatment that is applied.

But predators often are restricted to only certain microhabitats or substrate types, so that the presence or absence of the predator is indeed naturally confounded with differences in substrate type. This makes it difficult to tease apart cause and effect (see Chapter 6). The strength of multifactor field experiments is that they break apart this natural covariation and reveal the effects of multiple factors separately and in concert. The fact that some of these treatment combinations may be artificial and rarely, if ever, found in nature actually is a strength of the experiment: it reveals the independent contribution of each factor to the observed patterns.

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Figure 7.10 Example of a twoway design. Treatment symbols are given in Figure 7.9. The spreadsheet contains columns to indicate which substrate treatment and which predation treatment were applied to each replicate. The entire design includes 4 × 3 × 10 = 120 replicates total, but only 36 replicates (three per treatment combination) are illustrated.

ID number: 1 Substrate treatment: Granite Predation treatment: Unmanipulated Number of barnacle recruits: 12

ID number 1 2 3 4 5 6 7 8 9 . . 120

Substrate treatment Granite Slate Cement Granite Slate Cement Granite Slate Cement . . Cement

ID number: 120 Substrate treatment: Cement Predation treatment: Inclusion Number of barnacle recruits: 2 ID number: 5 Substrate treatment: Slate Predation treatment: Control Number of barnacle recruits: 10

Predation treatment Unmanipulated Unmanipulated Unmanipulated Control Control Control Predator exclusion Predator exclusion Predator exclusion . . Predator inclusion

Number of barnacle recruits 12 10 8 14 10 8 50 68 39 . . 2

The key advantage of two-way designs is the ability to tease apart main effects and interactions between two factors. As we will discuss in Chapter 10, the interaction term represents the non-additive component of the response. The interaction measures the extent to which different treatment combinations act additively, synergistically, or antagonistically. Perhaps the main disadvantage of the two-way design is that the number of treatment combinations can quickly become too large for adequate replication.

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In the barnacle predation example, 120 total replicates are required to replicate each treatment combination 10 times. As with the one-way layout, a simple two-way layout does not account for spatial heterogeneity. This can be handled by a simple randomized block design, in which every block contains one replicate each of all 12 treatment combinations. Alternatively, if you replicate all of the treatments within each block, this becomes a three-way design, with the blocks forming the third factor in the analysis. A final limitation of two-way designs is that it may not be possible to establish all orthogonal treatment combinations. It is somewhat surprising that for many common ecological experiments, the full set of treatment combinations may not be feasible or logical. For example, suppose you are studying the effects of competition between two species of salamanders on salamander survival rate. You decide to use a simple two-way design in which each species represents one of the factors. Within each factor, the two treatments are the presence or absence of the species. This fully crossed design yields four treatments (Table 7.2). But what are you going to measure in the treatment combination that has neither Species A nor Species B? By definition, there is nothing to measure in this treatment combination. Instead, you will have to establish the other three treatments ([Species A Present, Species B Absent], [Species A Absent, Species B Present], [Species A Present, Species B Present]) and analyze the design as a one-way ANOVA. The two-way design is possible only if we change the response variable. If the response variable is the abundance of salamander prey remaining

TABLE 7.2 Treatment combinations in a two-way layout for simple species addition and removal experiments Species B Absent Present

Species A Absent

Present

10 10

The entry in each cell is the number of replicates of each treatment combination. If the response variable is some property of the species themselves (e.g., survivorship, growth rate), then the treatment combination Species A Absent–Species B Absent (boxed) is not logically possible, and the analysis will have to use a one-way layout with three treatment groups (Species A Present–Species B Present, Species A Present–Species B Absent, and Species A Absent–Species B Present). If the response variable is some property of the environment that is potentially affected by the species (e.g., prey abundance, pH), then all four treatment combinations can be used and analyzed as a two-way ANOVA with two orthogonal factors (Species A and Species B), each with two treatment levels (Absent, Present).

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in each plot at the end of the experiment, rather than salamander survivorship, we can then establish the treatment with no salamanders of either species and measure prey levels in the fully crossed two-way layout. Of course, this experiment now asks an entirely different question. Two-species competition experiments like our salamander example have a long history in ecological and environmental research (Goldberg and Scheiner 2001). A number of subtle problems arise in the design and analysis of twospecies competition experiments. These experiments attempt to distinguish between a focal species, for which the response variable is measured, an associative species, whose density is manipulated, and background species, which may be present, but are not experimentally manipulated. The first issue is what kind of design to use: additive, substitutive, or response surface (Figure 7.11; Silvertown 1987). In an additive design, the density of the focal species is kept constant while the density of the experimental species is varied. However, this design confounds both density and frequency effects. For example, if we compare a control plot (5 individuals of Species A, 0 individuals of Species B) to an addition plot (5 individuals of Species A, 5 individuals of Species B), we have confounded total density (10 individuals) with the presence of the competitor (Underwood 1986; Bernardo et al. 1995). On the other hand, some

Substitutive design

Number of individuals of Species B

Response surface design ×

Additive design

0 2 4 1 Number of individuals of Species A

Figure 7.11 Experimental designs for competition experiments. The abundance of Species A and B are each set at 0, 1, 2, or 4 individuals. Each × indicates a different treatment combination. In an additive design, the abundance of one species is fixed (2 individuals of Species A) and the abundance of the competitor is varied (0, 1, 2, or 4 individuals of Species B). In a substitutive design, the total abundance of both competitors is held constant at 4 individuals, but the species composition in the different treatments is altered (0,4; 1,3; 2,2; 3,1; 4,0). In a response surface design, all abundance combinations of the two competitors are established in different treatments (4 × 4 = 16 treatments). The response surface design is preferred because it follows the principle of a good two-way ANOVA: the treatment levels are fully orthogonal (all abundance levels of Species A are represented with all abundance levels of Species B). (See Inouye 2001 for more details. Figure modified from Goldberg and Scheiner 2001.)

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authors have argued that such changes in density are indeed observed when a new species enters a community and establishes a population, so that adjusting for total density is not necessarily appropriate (Schluter 1995). In a substitutive design, total density of organisms is kept constant, but the relative proportions of the two competitors are varied. These designs measure the relative intensity of inter- and intraspecific competition, but they do not measure the absolute strength of competition, and they assume responses are comparable at different density levels. The response-surface design is a fully-crossed, two-way design that varies both the relative proportion and density of competitors. This design can be used to measure both relative intensity and absolute strength of inter- and intraspecific competitive interactions. However, as with all two-factor experiments with many treatment levels, adequate replication may be a problem. Inouye (2001) thoroughly reviews response-surface designs and other alternatives for competition studies. Other issues that need to be addressed in competition experiments include: how many density levels to incorporate in order to estimate accurately competitive effects; how to deal with the non-independence of individuals within a treatment replicate; whether to manipulate or control for background species; and how to deal with residual carry-over effects and spatial heterogeneity that are generated by removal experiments, in which plots are established based on the presence of a species (Goldberg and Scheiner 2001). SPLIT-PLOT DESIGNS

The split-plot design is an extension of the randomized block design to two experimental treatments. The terminology comes from agricultural studies in which a single plot is split into subplots, each of which receives a different treatment. For our purposes, such a split-plot is equivalent to a block that contains within it different treatment replicates. What distinguishes a split-plot design from a randomized block design is that a second treatment factor is also applied, this time at the level of the entire plot. Let’s return one last time to the barnacle example. Once again, you are going to set up a two-way design, testing for predation and substrate effects. However, suppose that the cages are expensive and time-consuming to construct, and that you suspect there is a lot of microhabitat variation in the environment that is affecting your results. In a split-plot design, you would group the three substrates together, just as you did in the randomized block design. However, you would then place a single cage over all three of the substrate replicates within a single block. In this design, the predation treatment is referred to as the wholeplot factor because a single predation treatment is applied to an entire block. The substrate treatment is referred to as the subplot factor because all substrate treatments are applied within a single block. The split-plot design is illustrated in Figure 7.12.

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Figure 7.12 Example of a splitplot design. Treatment symbols are given in Figure 7.9. The three substrate treatments (subplot factor) are grouped in blocks. The predation treatment (whole plot factor) is applied to an entire block. The spreadsheet contains columns to indicate the substrate treatment, predation treatment, and block identity for each replicate. Only a subset of the blocks in each predation treatment are illustrated. The split-plot design is similar to a randomized block design (see Figure 7.6), but in this case a second treatment factor is applied to the entire block (= plot).

ID number: 120 Substrate treatment: Cement Predation treatment: Inclusion Block number: 40 Number of barnacle recruits: 2 ID number 1 2 3 4 5 6 7 8 9 . . 120

Substrate treatment Granite Slate Cement Granite Slate Cement Granite Slate Cement . . Cement

ID number: 1 Substrate treatment: Granite Predation treatment: Unmanipulated Block number: 1 Number of barnacle recruits: 12

Predation treatment Unmanipulated Unmanipulated Unmanipulated Control Control Control Predator exclusion Predator exclusion Predator exclusion . . Predator inclusion

Block number 1 1 1 2 2 2 3 3 3 . . 40

Number of barnacle recruits 12 10 8 14 10 8 50 68 39 . . 2

You should compare carefully the two-way layout (Figure 7.10), and the splitplot layout (Figure 7.12) and appreciate the subtle difference between them. The distinction is that, in the two-way layout, each replicate receives the treatment applications independently and separately. In the split-plot layout, one of the treatments is applied to entire blocks or plots, and the other treatment is applied to replicates within blocks.

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The chief advantage of the split-plot design is the efficient use of blocks for the application of two treatments. As in the randomized block design, this is a simple layout that controls for environmental heterogeneity. It also may be less labor-intensive than applying treatments to individual replicates in a simple twoway design. The split-plot design removes the additive effects of the blocks and allows for tests of the main effects and interactions between the two manipulated factors.7 As in the randomized block design, the split-plot design does not allow you to test for the interaction between blocks and the subplot factor. However, the split-plot design does let you test for the main effect of the whole-plot factor, the main effect of the subplot factor, and the interaction between the two. As with nested designs, a very common mistake is for investigators to analyze a split-plot design as a two-factor ANOVA, which increases the risk of a Type I error. The two-way design can be extended to three or even more factors. For example, if you were studying trophic cascades in a freshwater food web (Brett and Goldman 1997), you might add or remove top carnivores, predators, and herbivores, and then measure the effects on the producer level. This simple three-way design generates 23 = 8 treatment combinations, including one combination that has neither top carnivores, predators, nor herbivores (Table 7.3). As we noted above, if you set up a randomized block design with a two-way layout and then replicate within blocks, the blocks then become a third factor in the analysis. However, three-factor (and higher) designs are used rarely in ecological studies. There are simply too many treatment combinations to make these designs logistically feasible. If you find

DESIGNS FOR THREE OR MORE FACTORS

Although the example we presented used two experimentally manipulated factors, the split-plot design is also effective when one of the two factors represents a source of natural variation. For example, in our research, we have studied the organization of aquatic food webs that develop in the rain-filled leaves of the pitcher plant Sarracenia purpurea. A new pitcher opens about once every 20 days, fills with rainwater, and quickly develops an associated food web of invertebrates and microorganisms. In one of our experiments, we manipulated the disturbance regime by adding or removing water from the leaves of each plant (Gotelli and Ellison 2006; see also Figure 9.15). These water manipulations were applied to all of the pitchers of a plant. Next, we recorded food web structure in the first, second, and third pitchers. These data are analyzed as a split-plot design. The whole-plot factor is water treatment (5 levels) and the subplot factor is pitcher age (3 levels). The plant served as a natural block, and it was efficient and realistic to apply the water treatments to all the leaves of a plant.

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TABLE 7.3 Treatment combinations in a three-way layout for a food web addition and removal experiment Carnivore absent

Producer absent Producer present

Carnivore present

Herbivore absent

Herbivore present

Herbivore absent

Herbivore present

10 10

In this experiment, the three trophic groups represent the three experimental factors (Carnivore, Herbivore, Producer), each of which has two levels (Absent, Present). The entry in each cell is the number of replicates of each treatment combination. If the response variable is some property of the food web itself, then the treatment combination in which all three trophic levels are absent (boxed) is not logically possible.

your design becoming too large and complex, you should consider breaking it down into a number of smaller experiments that address the key hypotheses you want to test. INCORPORATING TEMPORAL VARIABILITY: REPEATED MEASURES DESIGNS

In all of the designs we have described so far, the response variable is measured for each replicate at a single point in time at the end of the experiment. A repeated measures design is used whenever multiple observations on the same replicate are collected at different times. The repeated measures design can be thought of as a split-plot design in which a single replicate serves as a block, and the subplot factor is time. Repeated measures designs were first used in medical and psychological studies in which repeated observations were taken on an individual subject. Thus, in repeated measures terminology, the between-subjects factor corresponds to the whole-plot factor, and the within-subjects factor corresponds to the different times. In a repeated measures design, however, the multiple observations on a single individual are not independent of one another, and the analysis must proceed cautiously. For example, suppose we used the simple one-way design for the barnacle study shown in Figure 7.5. But rather than censusing each replicate once, we measured the number of new barnacle recruits on each replicate for 4 consecutive weeks. Now, instead of 3 treatments × 10 replicates = 30 observations, we have 3 treatments × 10 replicates × 4 weeks = 120 observations (Table 7.4). If we only used data from one of the four censuses, the analysis would be identical to the one-way layout. There are three advantages to a repeated measures design; the first is efficiency. Data are recorded at different times, but it is not necessary to have unique replicates for each time× treatment combination. Second, the repeated measures

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TABLE 7.4 Spreadsheet for a simple repeated measures analysis Barnacle recruits ID number

1 2 3 4 5 6 7 8 9 ... 30

Treatment

Granite Slate Cement Granite Slate Cement Granite Slate Cement ... Cement

Replicate

Week 1

Week 2

Week 3

Week 4

1 1 1 2 2 2 3 3 3 ... 10

12 10 3 14 10 8 11 11 7 ... 8

15 6 2 14 11 9 13 17 7 .. 0

17 19 0 5 13 4 22 28 7 ... 0

17 32 2 11 15 4 29 15 6 ... 3

This experiment is designed to test for the effect of substrate type on barnacle recruitment in the rocky intertidal (see Figure 7.5). The data are organized in a spreadsheet in which each row is an independent replicate. The columns indicate the ID number (1–30), the treatment group (Cement, Slate, or Granite), and the replicate number (1–10 within each treatment). The next four columns give the number of barnacle recruits recorded on a particular substrate in each of four consecutive weeks. The measurements at different times are not independent of one another because they are taken from the same replicate each week.

design allows each replicate to serve as its own block or control. When the replicates represent individuals (plants, animals, or humans), this effectively controls for variation in size, age, and individual history, which often have strong influences on the response variable. Finally, the repeated measures design allows us to test for interactions of time with treatment. For many reasons, we expect that differences among treatments may change with time. In a press experiment (see Chapter 6), there may be cumulative effects of the treatment that are not expressed until some time after the start of the experiment. In contrast, in a pulse experiment, we expect to see differences among treatments diminish as more time passes following the single, pulsed treatment application. Such complex effects are best seen in the interaction between time and treatment, and they may not be detected if the response variable is measured at only a single point in time. Both the randomized block and the repeated measures designs make a special assumption of circularity for the within-subjects factor. Circularity (in the context of ANOVA) means that the variance of the difference between any two treatment levels in the subplot is the same. For the randomized block design, this means that the variance of the difference between any pair of treat-

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ments in the block is the same. If the treatment plots are large enough and adequately spaced, this is often a reasonable assumption. For the repeated measures design, the assumption of circularity means that the variance of the difference of observations between any pair of times is the same. This assumption of circularity is unlikely to be met for repeated measures; in most cases, the variance of the difference between two consecutive observations is likely to be much smaller than the variance of the difference between two observations that are widely separated in time. This is because time series measured on the same subject are likely to have a temporal “memory” such that current values are a function of values observed in the recent past. This premise of correlated observations is the basis for time-series analysis (see Chapter 6). The chief disadvantage with repeated measures analysis is failure to meet the assumption of circularity. If the repeated measures are serially correlated, Type I error rates for F-tests will be inflated, and the null hypothesis may be incorrectly rejected when it is true. The best way to meet the circularity assumption is to use evenly spaced sampling times along with knowledge of the natural history of your organisms to select an appropriate sampling interval. What are some alternatives to repeated measures analysis that do not rely on the assumption of circularity? One approach is to set out enough replicates so that a different set is censused at each time period. With this design, time can be treated as a simple factor in a two-way analysis of variance. If the sampling methods are destructive (e.g., collecting stomach contents of fish, killing and preserving an invertebrate sample, or harvesting plants), this is the only method for incorporating time into the design. A second strategy is to use the repeated measures layout, but to be more creative in the design of the response variable. Collapse the correlated repeated measures into a single response variable for each individual, and then use a simple one-way analysis of variance. For example, if you want to test whether temporal trends differ among the treatments (the between-subjects factor), you could fit a regression line (with either a linear or time-series model) to the repeated measures data, and use the slope of the line as the response variable. A separate slope value would be calculated for each of the individuals in the study. The slopes would then be compared using a simple one-way analysis, treating each individual as an independent observation (which it is). Significant treatment effects would indicate different temporal trajectories for individuals in the different treatments; this test is very similar to the test for an interaction of time and treatment in a standard repeated measures analysis. Although such composite variables are created from correlated observations collected from one individual, they are independent among individuals. Moreover, the Central Limit Theorem (see Chapter 2) tells us that averages of

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these values will follow an approximately normal distribution even if the underlying variables themselves do not. Because most repeated measures data do not meet the assumption of circularity, our advice is to be careful with these analyses. We prefer to collapse the temporal data to a single variable that is truly independent among observations and then use a simpler one-way design for the analysis. ENVIRONMENTAL IMPACTS OVER TIME: BACI DESIGNS A special type of repeated measures design is one in which measurements are taken both before and after the application of a treatment. For example, suppose you are interested in measuring the effect of atrazine (a hormone-mimicking compound) on the body size of frogs (Allran and Karasov 2001). In a simple one-way layout, you could assign frogs randomly to control and treatment groups, apply the atrazine, and measure body size at the end of the experiment. A more sensitive design might be to establish the control and treatment groups, and take measurements of body size for one or more time periods before application of the treatment. After the treatment is applied, you again measure body size at several times in both the control and treatment groups. These designs also are used for observational studies assessing environmental impacts. In impact assessment, the measurements are taken before and after the impact occurs. A typical assessment might be the potential responses of a marine invertebrate community to the operation of a nuclear power plant, which discharges considerable hot water effluent (Schroeter et al. 1993). Before the power plant begins operation, you take one or more samples in the area that will be affected by the plant and estimate the abundance of species of interest (e.g., snails, sea stars, sea urchins). Replication in this study could be spatial, temporal, or both. Spatial replication would require sampling several different plots, both within and beyond the projected plume of hot water discharge.8 Temporal replication would require sampling a single site in the discharge area at several times before the plant came on-line. Ideally, multiple sites would be sampled several times in the pre-discharge period. Once the discharge begins, the sampling protocol is repeated. In this assessment design, it is imperative that there be at least one control or reference site that is sampled at the same time, both before and after the discharge. Then, if you observe a

A key assumption in this layout is that the investigator knows ahead of time the spatial extent of the impact. Without this information, some of the “control” plots may end up within the “treatment” region, and the effect of the hot water plume would be underestimated.

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decline in abundance at the impacted site but not at the control site, you can test whether the decline is significant. Alternatively, invertebrate abundance might be declining for reasons that have nothing to do with hot water discharge. In this case, you would find lower abundance at both the control and the impacted sites. This kind of repeated measures design is referred to as a BACI design (BeforeAfter, Control-Impact). Not only is there replication of control and treatment plots, there is temporal replication with measurements before and after treatment application (Figure 7.13). In its ideal form, the BACI design is a powerful layout for assessing environmental perturbations and monitoring trajectories before and after the impact. Replication in space ensures that the results will be applicable to other sites that may be perturbed in the same way. Replication through time ensures that the temporal trajectory of response and recovery can be monitored. The design is appropriate for both pulse and press experiments or disturbances. Unfortunately, this idealized BACI design is rarely achieved in environmental impact studies. Often times, there is only a single site that will be impacted, and that site usually is not chosen randomly (Stewart-Oaten and Bence 2001; Murtaugh 2002b). Spatial replicates within the impacted area are not independent replicates because there is only a single impact studied at a single site (Underwood 1994). If the impact represents an environmental accident, such as an oil spill, no pre-impact data may be available, either from reference or impact sites. The potential control of randomization and treatment assignments is much better in large-scale experimental manipulations, but even in these cases there may be little, if any, spatial replication. These studies rely on more intensive temporal replication, both before and after the manipulation. For example, since 1983, Brezonik et al. (1986) have conducted a long-term acidification experiment on Little Rock Lake, a small oligotrophic seepage lake in northern Wisconsin. The lake was divided into a treatment and reference basin with an impermeable vinyl curtain. Baseline (pre-manipulation) data were collected in both basins from August 1983 through April 1985. The treatment basin was then acidified with sulfuric acid in a stepwise fashion to three target pH levels (5.6, 5.1, 4.7). These pH levels were maintained in a press experiment at two-year intervals. Because there is only one treatment basin and one control basin, conventional ANOVA methods cannot be used to analyze these data.9 There are two general 9

Of course, if one assumes the samples are independent replicates, conventional ANOVA could be used. But there is no wisdom in forcing data into a model structure they do not fit. One of the key themes of this chapter is to choose simple designs for your experiments and surveys whose assumptions best meet the constraints of your data.

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Effluent discharge impact zone

ID number: 1 Treatment: Control Power plant

ID number 1 2 3 4 5 6 7 8

ID number: 6 Treatment: Impact

Pre-impact sampling Post-impact sampling Treatment Week 1 Week 2 Week 3 Week 4 Week 5 Week 6 Week 7 Week 8 Control 106 190 130 123 122 120 108 108 Control 104 120 135 119 106 104 84 88 Impact 99 150 145 140 150 192 102 97 Impact 120 165 155 135 137 120 98 122 77 75 7 0 94 92 90 Impact 88 Impact 100 130 66 3 2 82 129 120 109 55 55 45 99 70 70 Control 66 140 88 90 100 250 220 209 Control 130

Figure 7.13 Example of a spatial arrangement of replicates for a BACI design. Each square represents a sample plot on a shoreline that will potentially be affected by hot water discharged from a nuclear power plant. Permanent plots are established within the hot water effluent zone (shaded area), and in adjacent control zones (unshaded areas). All plots are sampled weekly for 4 weeks before the plant begins discharging hot water and for 4 weeks afterward. Each row of the spreadsheet represents a different replicate. Two columns indicate the replicate ID number and the treatment (Control or Impact). The remaining columns give the invertebrate abundance data collected at each of the 8 sampling dates (4 sample dates pre-discharge, 4 sample dates post-discharge).

strategies for analysis. Randomized intervention analysis (RIA) is a Monte Carlo procedure (see Chapter 5) in which a test statistic calculated from the time series is compared to a distribution of values created by randomizing or reshuffling the time-series data among the treatment intervals (Carpenter et al. 1989). RIA relaxes the assumption of normality, but it is still susceptible to temporal correlations in the data (Stewart-Oaten et al. 1992). A second strategy is to use time-series analysis to fit simple models to the data. The autoregressive integrated moving average (ARIMA) model describes the correlation structure in temporal data with a few parameters (see Chapter 6). Additional model parameters estimate the stepwise changes that occur with the experimental interventions, and these parameters can then be tested against the

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null hypothesis that they do not differ from 0. ARIMA models can be fit individually to the control and manipulation time series data, or to a derived data series created by taking the ratio of the treatment/control data at each time step (Rasmussen et al. 1993). Bayesian methods can also be used to analyze data from BACI designs (Carpenter et al. 1996; Rao and Tirtotjondro 1996; Reckhow 1996; Varis and Kuikka 1997; Fox 2001). RIA, ARIMA, and Bayesian methods are powerful tools for detecting treatment effects in time-series data. However, without replication, it is still problematic to generalize the results of the analysis. What might happen in other lakes? Other years? Additional information, including the results of small-scale experiments (Frost et al. 1988), or snapshot comparisons with a large number of unmanipulated control sites (Schindler et al. 1985; Underwood 1994) can help to expand the domain of inference from BACI study. Alternatives to ANOVA: Experimental Regression

The literature on experimental design is dominated by ANOVA layouts, and modern ecological science has been referred to sarcastically as little more than the care and curation of ANOVA tables. Although ANOVA designs are convenient and powerful for many purposes, they are not always the best choice. ANOVA has become so popular that it may act as an intellectual straightjacket (Werner 1998), and cause scientists to neglect other useful experimental designs. We suggest that ANOVA designs often are employed when a regression design would be more appropriate. In many ANOVA designs, a continuous predictor variable is tested at only a few values so that it can be treated as a categorical predictor variable, and shoehorned into an ANOVA design. Examples include treatment levels that represent different nutrient concentrations, temperatures, or resource levels. In contrast, an experimental regression design (Figure 7.14) uses many different levels of the continuous independent variable, and then uses regression to fit a line, curve, or surface to the data. One tricky issue in such a design (the same problem is present in an ANOVA) is to choose appropriate levels for the predictor variable. A uniform selection of predictor values within the desired range should ensure high statistical power and a reasonable fit to the regression line. However, if the response is expected to be multiplicative rather than linear (e.g., a 10% decrease in growth with every doubling of concentration), it might be better to set the predictor values on an evenly spaced logarithmic scale. In this design, you will have more data collected at low concentrations, where changes in the response variable might be expected to be steepest. One of the chief advantages of regression designs is efficiency. Suppose you are studying responses of terrestrial plant and insect communities to nitrogen

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(Simultaneous N and P treatments in a two-way ANOVA design) 0.00 mg Phosphorus treatment 0.05 mg

(Two-way experimental regression design) 0.00 mg 0.00 mg 1 0.01 mg 1 0.05 mg 1 Phosphorus 0.10 mg 1 treatment 1 0.20 mg 1 0.40 mg 1 0.50 mg

0.05 mg 1 1 1 1 1 1 1

Nitrogen treatment 0.00 mg 0.50 mg 12 12 12 12

Nitrogen treatment 0.10 mg 0.20 mg 0.40 mg 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1

0.80 mg 1.00 mg 1 1 1 1 1 1 1 1 1 1 1 1 1 1

Figure 7.14 Treatment combinations in a two-way ANOVA design (upper panel) and an experimental regression design (lower panel). These experiments test for additive and interactive effects of nitrogen (N) and phosphorus (P) on plant growth or some other response variable. Each cell in the table indicates the number of replicate plots used. If the maximum number of replicates is 50 and a minimum of 10 replicates per treatment in required, only 2 treatment levels each for N and P are possible (each with 12 replicates) in the two-way ANOVA. In contrast, the experimental regression allows for 7 treatment levels each of N and P. Each of the 7 × 7 = 49 plots in the design receives a unique combination of N and P concentrations.

(N), and your total sample size is limited by available space or labor to 50 plots. If you try to follow the Rule of 10, an ANOVA design would force you to select only 5 different fertilization levels and replicate each one 10 times. Although this design is adequate for some purposes, it may not help to pinpoint critical threshold levels at which community structure changes dramatically in response to N. In contrast, a regression design would allow you to set up 50 different N levels, one in each of the plots. With this design, you could very accurately characterize changes that occur in community structure with increasing N levels; graphical displays may help to reveal threshold points and non-linear effects (see Chapter 9). Of course, even minimal replication of each treatment level is very desirable, but if the total sample size is limited, this may not be possible. For a two-factor ANOVA, the experimental regression is even more efficient and powerful. If you want to manipulate nitrogen and phosphorus (P) as independent factors, and still maintain 10 replicates per treatment, you could have no more than 2 levels of N and 2 levels of P. Because one of those levels must be a control plot (i.e., no fertilization), the experiment isn’t going to give you very much information about the role of changing levels of N and P on the

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system. If the result is statistically significant, you can say only that the community responds to those particular levels of N and P, which is something that you may have already known from the literature before you started. If the result is not statistically significant, the obvious criticism would be that the concentrations of N and P were too low to generate a response. In contrast, an experimental regression design would be a fully crossed design with 7 levels of nitrogen and 7 levels of phosphorus, with one level of each corresponding to the control (no N or P). The 7 × 7 = 49 replicates each receive a unique concentration of N and P, with one of the 49 plots receiving neither N nor P (see Figure 7.14). This is a response surface design (Inouye 2001), in which the response variable will be modeled by multiple regression. With seven levels of each nutrient, this design provides a much more powerful test for additive and interactive effects of nutrients, and could also reveal non-linear responses. If the effects of N and P are weak or subtle, the regression model will be more likely to reveal significant effects than the two-way ANOVA.10 Efficiency is not the only advantage of an experimental regression design. By representing the predictor variable naturally on a continuous scale, it is much easier to detect non-linear, threshold, or asymptotic responses. These cannot be inferred reliably from an ANOVA design, which usually will not have enough treatment levels to be informative. If the relationship is something other than a straight line, there are a number of statistical methods for fitting non-linear responses (see Chapter 9). A final advantage to using an experimental regression design is the potential benefits for integrating your results with theoretical predictions and ecological models. ANOVA provides estimates of means and variances for groups or particular levels of categorical variables. These estimates are rarely of interest or use in ecological models. In contrast, a regression analysis provides esti-

There is a further hidden penalty in using the two-way ANOVA design for this experiment that often is not appreciated. If the treatment levels represent a small subset of many possible other levels that could have been used, then the design is referred to as a random effects ANOVA model. Unless there is something special about the particular treatment levels that were used, a random effects model is always the most appropriate choice when a continuous variable has been shoehorned into a categorical variable for ANOVA. In the random effects model, the denominator of the F-ratio test for treatment effects is the interaction mean square, not the error mean square that is used in a standard fixed effects ANOVA model. If there are not many treatment levels, there will not be very many degrees of freedom associated with the interaction term, regardless of the amount of replication within treatments. As a consequence, the test will be much less powerful than a typical fixed effects ANOVA. See Chapter 10 for more details on fixed and random effects ANOVA models and the construction of F-ratios.

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mates of slope and intercept parameters that measure the change in the response Y relative to a change in predictor X (dY/dX). These derivatives are precisely what are needed for testing the many ecological models that are written as simple differential equations. An experimental regression approach might not be feasible if it is very expensive or time-consuming to establish unique levels of the predictor variable. In that case, an ANOVA design may be preferred because only a few levels of the predictor variable can be established. An apparent disadvantage of the experimental regression is that it appears to have no replication! In Figure 7.14, each unique treatment level is applied to only a single replicate, and that seems to fly in the face of the principle of replication (see Chapter 6). If each unique treatment is unreplicated, the least-squares solution to the regression line still provides an estimate of the regression parameters and their variances (see Chapter 9). The regression line provides an unbiased estimate of the expected value of the response Y for a given value of the predictor X, and the variance estimates can be used to construct a confidence interval about that expectation. This is actually more informative than the results of an ANOVA model, which allow you to estimate means and confidence intervals for only the handful of treatment levels that were used. A potentially more serious issue is that the regression design may not include any replication of controls. In our two-factor example, there is only one plot that contains no nitrogen and no phosphorus addition. Whether this is a serious problem or not depends on the details of the experimental design. As long as all of the replicates are treated the same and differ only in the treatment application, the experiment is still a valid one, and the results will estimate accurately the relative effects of different levels of the predictor variable on the response variable. If it is desirable to estimate the absolute treatment effect, then additional replicated control plots may be needed to account for any handling effects or other responses to the general experimental conditions. These issues are no different than those that are encountered in ANOVA designs. Historically, regression has been used predominantly in the analysis of nonexperimental data, even though its assumptions are unlikely to be met in most sampling studies. An experimental study based on a regression design not only meets the assumptions of the analysis, but is often more powerful and appropriate than an ANOVA design. We encourage you to “think outside the ANOVA box” and consider a regression design when you are manipulating a continuous predictor variable. Tabular Designs

The last class of experimental designs is used when both predictor and response variables are categorical. The measurements in these designs are counts. The

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Four Classes of Experimental Design

simplest such variable is a dichotomous (or binomial, see Chapter 2) response in a series of independent trials. For example, in a test of co*ckroach behavior, you could place an individual co*ckroach in an arena with a black and a white side, and then record on which side the animal spent the majority of its time. To ensure independence, each replicate co*ckroach would be tested individually. More typically, a dichotomous response will be recorded for two or more categories of the predictor variable. In the co*ckroach study, half of the co*ckroaches might be infected experimentally with a parasite that is known to alter host behavior (Moore 1984). Now we want to ask whether the response of the co*ckroach differs between parasitized and unparasitized individuals (Moore 2001; Poulin 2000). This approach could be extended to a three-way design by adding an additional treatment and asking whether the difference between parasitized and unparasitized individuals changes in the presence or absence of a vertebrate predator. We might predict that uninfected individuals are more likely to use the black substrate, which will make them less conspicuous to a visual predator. In the presence of a predator, uninfected individuals might shift even more toward the dark surfaces, whereas infected individuals might shift more toward white surfaces. Alternatively, the parasite might alter host behavior, but those alterations might be independent of the presence or absence of the predator. Still another possibility is that host behavior might be very sensitive to the presence of the predator, but not necessarily affected by parasite infection. A contingency table analysis (see Chapter 11) is used to test all these hypotheses with the same dataset. In some tabular designs, the investigator determines the total number of individuals in each category of predictor variable, and these individuals will be classified according to their responses. The total for each category is referred to as the marginal total because it represents the column or row sum in the margin of the data table. In an observational study, the investigator might determine one or both of the marginal totals, or perhaps only the grand total of independent observations. In a tabular design, the grand total equals the sum of either the column or row marginal totals. For example, suppose you are trying to determine the associations of four species of Anolis lizard with three microhabitat types (ground, tree trunks, tree branches; see Butler and Losos 2002). Table 7.5 shows the two-way layout of the data from such a study. Each row in the table represents a different lizard species, and each column represents a different habitat category. The entries in each cell represent the counts of a particular lizard species recorded in a particular habitat. The marginal row totals represent the total number of observations for each lizard species, summed across the three habitat types. The marginal column totals

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TABLE 7.5 Tabulated counts of the occurrence of four lizard species censused in three different microhabitats Habitat

Lizard species

Species A Species B Species C Species D Habitat totals

Ground

Tree trunk

Tree branch

Species totals

9 9 9 9

0 0 5 10

15 12 0 3

24 21 14 22

Italicized values are the marginal totals for the two-way table. The total sample size is 81 observations. In these data, both the response variable (species identity) and the predictor variable (microhabitat category) are categorical.

represent the total number of observations in each habitat type, summed across the three habitats. The grand total in the table (N = 81) represents the total count of all lizard species observed in all habitats. There are several ways that these data could have been collected, depending on whether the sampling was based on the marginal totals for the microhabitats, the marginal totals for the lizards, or the grand total for the entire sample. In a sampling scheme built around the microhabitats, the investigator might have spent 10 hours sampling each microhabitat, and recording the number of different lizard species encountered in each habitat. Thus, in the census of tree trunks, the investigator found a total of 15 lizards: 5 of Species C and 10 of Species D; Species A and B were never encountered. In the ground census, the investigator found a total of 36 lizards, with all four species equally represented. Alternatively, the sampling could have been based on the lizards themselves. In such a design, the investigator would put in an equal sampling effort for each species by searching all habitats randomly for individuals of a particular species of lizard and then recording in which microhabitat they occurred. Thus, in a search for Species B, 21 individuals were found, 9 on the ground, and 12 on tree branches. Another sampling variant is one in which the row and column totals are simultaneously fixed. Although this design is not very common in ecological studies, it can be used for an exact statistical test of the distribution of sampled values (Fisher’s Exact Test; see Chapter 11). Finally, the sampling might have been based simply on the grand total of observations. Thus, the investigator might have taken a random sample of 81 lizards, and for each lizard encountered, recorded the species identity and the microhabitat. Ideally, the marginal totals on which the sampling is based should be the same for each category, just as we try to achieve equal sample sizes in setting up an

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Four Classes of Experimental Design

ANOVA design. However, identical sample sizes are not necessary for analysis of tabular data. Nevertheless, the tests do require (as always) that the observations be randomly sampled and that the replicates be truly independent of one another. This may be very difficult to achieve in some cases. For example, if lizards tend to aggregate or move in groups, we cannot simply count individuals as they are encountered because an entire group is likely to be found in a single microhabitat. In this example, we also assume that all of the lizards are equally conspicuous to the observer in all of the habitats. If some species are more obvious in certain habitats than others, then the relative frequencies will reflect sampling biases rather than species’ microhabitat associations. SAMPLING DESIGNS FOR CATEGORICAL DATA In contrast to the large literature for regression and ANOVA designs, relatively little has been written, in an ecological context, about sampling designs for categorical data. If the observations are expensive or time-consuming, every effort should be made to ensure that each observation is independent, so that a simple two- or multiway layout can be used. Unfortunately, many published analyses of categorical data are based on nonindependent observations, some of it collected in different times or in different places. Many behavioral studies analyze multiple observations of the same individual. Such data clearly should not be treated as independent (Kramer and Schmidhammer 1992). If the tabular data are not independent, random samples from the same sampling space, you should explicitly incorporate the temporal or spatial categories as factors in your analysis.

Alternatives to Tabular Designs: Proportional Designs

If the individual observations are inexpensive and can be gathered in large numbers, there is an alternative to tabular designs. One of the categorical variables can be collapsed to a measure of proportion (number of desired outcomes/number of observations), which is a continuous variable. The continuous variable can then be analyzed using any of the methods described above for regression or ANOVA. There are two advantages of using proportional designs in lieu of tabular ones. The first is that the standard set of ANOVA and regression designs can be used, including blocking. The second advantage is that the analysis of proportions can be used to accommodate frequency data that are not strictly independent. For example, suppose that, to save time, the co*ckroach experiment were set up with 10 individuals placed in the behavioral arena at one time. It would not be legitimate to treat the 10 individuals as independent replicates, for the same reason that subsamples from a single cage are not independent replicates for an ANOVA. However, the data from this run can be treated as a single repli-

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cate, for which we could calculate the proportion of individuals present on the black side of the arena. With multiple runs of the experiment, we can now test hypotheses about differences in the proportion among groups (e.g., parasitized versus unparasitized). The design is still problematic, because it is possible that substrate selection by solitary co*ckroaches may be different from substrate selection by groups of co*ckroaches. Nevertheless, the design at least avoids treating individuals within an arena as independent replicates, which they are not. Although proportions, like probabilities, are continuous variables, they are bounded between 0.0 and 1.0. An arcsin square root transformation of proportional data may be necessary to meet the assumption of normality (see Chapter 8). A second consideration in the analysis of proportions is that it is very important to use at least 10 trials per replicate, and to make sure that sample sizes are as closely balanced as possible. With 10 individuals per replicate, the possible measures for the response variable are in the set {0.0, 0.1, 0.2, 0.3, 0.4, 0.5, 0.6, 0.7, 0.8, 0.9, 1.0}. But suppose the same treatment is applied to a replicate in which only three individuals were used. In this case, the only possible values are in the set {0.0, 0.33, 0.66, 1.0}. These small sample sizes will greatly inflate the measured variance, and this problem is not alleviated by any data transformation. A final problem with the analysis of proportions arises if there are three or more categorical variables. With a dichotomous response, the proportion completely characterizes the data. However, if there are more than two categories, the proportion will have to be carefully defined in terms of only one of the categories. For example, if the arena includes vertical and horizontal black and white surfaces, there are now four categories from which the proportion can be measured. Thus, the analysis might be based on the proportion of individuals using the horizontal black surface. Alternatively, the proportion could be defined in terms of two or more summed categories, such as the proportion of individuals using any vertical surface (white or black).

Summary Independent and dependent variables are either categorical or continuous, and most designs fit into one of four possible categories based on this classification. Analysis of variance (ANOVA) designs are used for experiments in which the independent variable is categorical and the dependent variable is continuous. Useful ANOVA designs include one- and two-way ANOVAs, randomized block, and split-plot designs. We do not favor the use of nested ANOVAs, in which non-independent subsamples are taken from within a replicate. Repeated measures designs can be used when repeated observations are collected on a single replicate through time. However, these data are often autocorrelated, so

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Summary

that the assumptions of the analysis may not be met. In such cases, the temporal data should be collapsed to a single independent measurement, or time-series analysis should be employed. If the independent variable is continuous, a regression design is used. Regression designs are appropriate for both experimental and sampling studies, although they are used predominantly in the latter. We advocate increased use of experimental regression in lieu of ANOVA with only a few levels of the independent variable represented. Adequate sampling of the range of predictor values is important in designing a sound regression experiment. Multiple regression designs include two or more predictor variables, although the analysis becomes problematic if there are strong correlations (collinearity) among the predictor variables. If both the independent and the dependent variables are categorical, a tabular design is employed. Tabular designs require true independence of the replicate counts. If the counts are not independent, they should be collapsed so that the response variable is a single proportion. The experimental design is then similar to a regression or ANOVA. We favor simple experimental and sampling designs and emphasize the importance of collecting data from replicates that are independent of one another. Good replication and balanced sample sizes will improve the power and reliability of the analyses. Even the most sophisticated analysis cannot salvage results from a poorly designed study.

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CHAPTER 8

Managing and Curating Data

Data are the raw material of scientific studies. However, we rarely devote the same amount of time and energy to data organization, management, and curation that we devote to data collection, analysis, and publication. This is an unfortunate state of affairs; it is far easier to use the original (raw) data themselves to test new hypotheses than it is to reconstruct the original data from summary tables and figures presented in the published literature. A basic requirement of any scientific study is that it be repeatable by others. Analyses cannot be reconstructed unless the original data are properly organized and documented, safely stored, and made available. Moreover, it is a legal requirement that any data collected as part of a project that is supported by public funds (e.g., federal or state grants and contracts) be made available to other scientists and to the public. Data sharing is legally mandated because the granting agency, not the investigator, technically owns the data. Most granting agencies allow ample time (usually one or more years) for investigators to publish their results before the data have to be made available publicly.1 However, the Freedom of Information Act (5 U.S.C. § 552) allows anyone in the United States to request publicly funded data from scientists at any time. For all of these reasons, we encourage you to manage your data as carefully as you collect, analyze, and publish them. 1

This policy is part of an unwritten gentleman’s agreement between granting agencies and the scientific community. This agreement works because the scientific community so far has demonstrated its willingness to make data available freely and in a timely fashion. Ecologists and environmental scientists generally are slower to make data available than are scientists working in more market-driven fields (such as biotechnology and molecular genetics) where there is greater public scrutiny. Similarly, provisions for data access and standards for metadata production by the ecological and environmental community have lagged behind those of other fields. It could be argued that the lack of a GenBank for ecologists has slowed the progress of ecological and environmental sciences. However, ecological data are much more heterogeneous than gene sequences, so it has proven difficult to design a single data structure to facilitate rapid data sharing.

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The First Step: Managing Raw Data In the laboratory-based bench sciences, the bound lab notebook is the traditional medium for recording observations. In contrast, ecological and environmental data are recorded in many forms. Examples include sets of observations written in notebooks, scrawled on plastic diving slates or waterproof paper, or dictated into voice recorders; digital outputs of instruments streamed directly into data loggers, computers, or palm-pilots; and silver-oxide negatives or digital satellite images. Our first responsibility with our data, therefore, is to transfer them rapidly from their heterogeneous origins into a common format that can be organized, checked, analyzed, and shared. Spreadsheets

In Chapter 5, we illustrated how to organize data in a spreadsheet. A spreadsheet is an electronic page2 in which each row represents a single observation (e.g., the unit of study, such as an individual leaf, feather, organism, or island), and each column represents a single measured or observed variable. The entry in each cell of the spreadsheet, or row-by-column pair, is the value for the unit of study (the row) of the measurement or observation of the variable (the column).3 All types of data used by ecologists and environmental scientists can be stored in spreadsheets, and the majority of experimental designs (see Chapter 7) can be represented in spreadsheet formats. Field data should be entered into spreadsheets as soon as possible after collection. In our research, we try to transfer the data from our field notes or instruments into spreadsheets on the same day that we collect it. There are several important reasons to transcribe your data promptly. First, you can determine quickly if necessary observations or experimental units were missed due to researcher oversight or instrument failure. If it is necessary to go back and col2

Although commercial spreadsheet software can facilitate data organization and management, it is not appropriate to use commercial software for the archival electronic copy of your data. Rather, archived data should be stored as ASCII (American Standard Code for Information Exchange) text files, which can be read by any machine without recourse to commercial packages that may not exist in 50 years. 3

We illustrate and work only with two-dimensional (row, column) spreadsheets (which are a type of flat files). Regardless of whether data are stored in two- or three-dimensional formats, pages within electronic workbooks must be exported as flat files for analysis with statistical software. Although spreadsheet software often contains statistical routines and random number generators, we do not recommend the use of these programs for analysis of scientific data. Spreadsheets are fine for generating simple summary statistics, but their random number generators and statistical algorithms may not be reliable for more complex analyses (Knüsel 1998; McCullough and Wilson 1999).

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The First Step: Managing Raw Data

lect additional observations, this must be done very quickly or the new data won’t be comparable. Second, your data sheets usually will include marginal notes and observations, in addition to the numbers you record. These marginal notes make sense when you take them, but they have a short half-life and rapidly become incomprehensible (and sometimes illegible). Third, once data are entered into a spreadsheet, you then have two copies of them; it is less likely you will misplace or lose both copies of your valuable results. Finally, rapid organization of data may promote timely analysis and publication; unorganized raw data certainly will not. Data should be proofread as soon as possible after entry into a spreadsheet. However, proofreading usually does not catch all errors in the data, and once the data have been organized and documented, further checking of the data is necessary (discussed later in the chapter). Metadata

At the same time that you enter the data into a spreadsheet, you should begin to construct the metadata that must accompany the data. Metadata are “data about data” and describe key attributes of the dataset. Minimal metadata4 that should accompany a dataset include: • • • • •

Name and contact information for the person who collected the data Geographic information about where the data were collected The name of the study for which the data were collected The source of support that enabled the data to be collected A description of the organization of the data file, which should include: • A brief description of the methods used to collect the data • The types of experimental units • The units of measurement or observation of each of the variables • A description of any abbreviations used in the data file • An explicit description of what data are in columns, what data are in rows, and what character is used to separate elements within rows

Three “levels” of metadata have been described by Michener et al. (1997) and Michener (2000). Level I metadata include a basic description of the dataset and the structure of the data. Level II metadata include not only descriptions of the dataset and its structure, but also information on the origin and location of the research, information on the version of the data, and instructions on how to access the data. Finally, Level III metadata are considered auditable and publishable. Level III metadata include all Level II metadata plus descriptions of how data were acquired, documentation of quality assurance and quality control (QA/QC), a full description of any software used to process the data, a clear description of how the data have been archived, a set of publications associated with the data, and a history of how the dataset has been used.

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(e.g., tabs, spaces, or commas) and between columns (e.g., a “hard return”).5 An alternative approach—more common in the laboratory sciences—is to create the metadata before the data are ever collected. A great deal of clear thinking can accompany the formal laying out and writing up of the methods, including plot design, intended number of observations, sample space, and organization of the data file. Such a priori metadata construction also facilitates the writing of the Methods section of the final report or publication of the study. We also encourage you to draw blank plots and tables that illustrate the relationships among variables that you propose to examine. Regardless of whether you choose to write up your metadata before or after conducting your study, we cannot overemphasize their importance. Even though it is time-consuming to assemble metadata, and they would seem to contain information that is self-evident, we guarantee that you will find them invaluable. You may think that you will never forget that Nemo Swamp was where you collected the data for your opus on endangered orchids. And of course you can still recall all the back roads you drove to get there. But time will erode those memories as the months and years pass, and you move on to new projects and studies. We all have experiences looking through lists of computer file names (NEMO, NEMO03, NEMO03NEW), trying to find the one we need. Even if we can find the needed data file, we may still be out of luck. Without metadata, there is no way to recollect the meaning of the alphabet soup of abbreviations and acronyms running across the top row of the file. Reconstructing datasets is frustrating, inefficient, and time-consuming, and impedes further data analysis. Undocumented datasets are nearly useless to you and to anyone else. Data without metadata cannot be stored in publicly-accessible data archives, and have no life beyond the summary statistics that describe them in a publication or report.

The Second Step: Storing and Curating the Data Storage: Temporary and Archival

Once the data are organized and documented, the dataset—a combination of data and metadata—should be stored on permanent media. The most perma5

Ecologists and environmental scientists continue to develop standards for metadata. These include the Content Standards for Digital Geospatial Metadata (FGDC 1998) for spatially-organized data (e.g., data from Geographic Information Systems, or GIS); standard descriptors for ecological and environmental data on soils (Boone et al. 1999); and Classification and Information Standards for data describing types of vegetation (FGDC 1997). Many of these have been incorporated within the Ecological Metadata Language, or EML, developed in late 2002 (knb.ecoinformatics.org/software/eml/) and which is now the de facto standard for documenting ecological data.

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The Second Step: Storing and Curating the Data

nent medium, and the only medium acceptable as truly archival, is acid-free paper. It is good practice to print out a copy of the raw data spreadsheet and its accompanying metadata onto acid-free paper using a laser printer.6 This copy should be stored somewhere that is safe from damage. Electronic storage of the original dataset also is a good idea, but you should not expect electronic media to last more than 5–10 years.7 Thus, electronic storage should be used primarily for working copies. If you intend the electronic copies to last more than a few years, you will have to copy the datasets onto newer electronic media on a regular basis. You must recognize that data storage has real costs, in space (where the data are stored), time (to maintain and transfer datasets among media), and money (because time and space are money). Curating the Data

Most ecological and environmental data are collected by researchers using funds obtained through grants and contracts. Usually, these data technically are owned by the granting agency, and they must be made available to others within a reasonably short period of time. Thus, regardless of how they are stored, your datasets must be maintained in a state that makes them usable not only by you, but also by the broader scientific community and other interested individuals and parties. Like museum and herbarium collections of animals and plants, datasets must be curated so they are accessible to others. Multiple datasets gen-

6 Inks from antique dot-matrix printers (and typewriters) last longer than those of inkjet printers, but the electrostatic bonding of ink to paper by laser printers lasts longer than either. True Luddites prefer Higgins Aeterna India Ink (with a tip o’ the hat to Henry Horn). 7 Few readers will remember floppy disks of any size (8”, 5-1/4”, 3-1/2”), much less paper tapes, punch cards (illustrated), or magnetic tape reels. When we wrote the first edition, CD-ROMs were being replaced rapidly by DVDROMs; in the intervening years, we have moved on to Computer punch card terabyte bricks, RAID arrays, and the seemingly ubiquitous “cloud.” Although the actual lifespan of most magnetic media is on the order of years to decades, and that of most optical media is on the order of decades-to-centuries (only years in the humid tropics where fungi eat the glue between the laminated layers of material and etch the disk), the capitalist marketplace replaces them all on an approximately 5-year cycle. Thus, although we still have on our shelves readable 5-1/4” floppy disks and magnetic tapes with data from the 1980s and 1990s, it is nearly impossible now to find, much less buy, a disk drive with which to read them. And even if we could find one, the current operating systems of our computers wouldn’t recognize it as usable hardware. A good resource for obsolete equipment and software is the Computer History Museum in Mountain View, California: www.computerhistory.org/.

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erated by a project or research group can be organized into electronic data catalogs. These catalogs often can be accessed and searched from remote locations using the internet. Nearly a decade after we wrote the first edition of this Primer, standards for curation of ecological and environmental data are still being developed. Most data catalogs are maintained on servers connected to the World Wide Web,8 and can be found using search engines that are available widely. Like data storage, data curation is costly; it has been estimated that data management and curation should account for approximately 10% of the total cost of a research project (Michener and Haddad 1992). Unfortunately, although granting agencies require that data be made available, they have been reluctant to fund the full costs of data management and curation. Researchers themselves rarely budget these costs in their grant proposals. When budgets inevitably are cut, data management and curation costs often are the first items to be dropped.

The Third Step: Checking the Data Before beginning to analyze a dataset, you must carefully examine it for outliers and errors. The Importance of Outliers

Outliers are recorded values of measurements or observations that are outside the range of the bulk of the data (Figure 8.1). Although there is no standard level of “outside the range” that defines an outlier, it is common practice to consider 8

Many scientists consider posting data on the World Wide Web or in the “cloud” to be a means of permanently archiving data. This is illusory. First, it is simply a transfer of responsibility from you to a computer system manager (or other information technology professional). By placing your electronic archival copy on the Web, you imply a belief that regular backups are made and maintained by the system manager. Every time a server is upgraded, the data have to be copied from the old server to the new one. Most laboratories or departments do not have their own systems managers, and the interests of computing centers in archiving and maintaining Web pages and data files do not necessarily parallel those of individual investigators. Second, server hard disks fail regularly (and often spectacularly). Last, the Web is neither permanent nor stable. GOPHER and LYNX have disappeared, FTP has all but been replaced by HTTP, and HTML, the original language of the Web, is steadily being replaced by (the not entirely compatible) XML. All of these changes to the functionality of the World Wide Web and the accessibility of files stored occurred within 10 years. It often is easier to recover data from notebooks that were hand-written in the nineteenth century than it is to recover data from Web sites that were digitally “archived” in the 1990s! In fact, it cost us more than $2000 in 2006 to recover from a 1997 magnetic tape the data used to illustrate cluster analysis and redundancy analysis in Chapter 12!

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The Third Step: Checking the Data

BC * *

A *

Number of plants

8 6 4 2 0 0

200

400

600 800 1000 Plant height (mm)

1200

1400

1600

Figure 8.1 Histogram and box plot of measurements of plant height for samples of the carnivorous pitcher plant Darlingtonia californica (sample size N = 25; data from Table 8.1). The histogram shows the frequency of plants in size class intervals of 100 mm. Above the histogram is the box plot of the same data. The vertical line in the box indicates the median of the distribution. The box encompasses 50% of the data, and the whiskers encompass 90% of the data. A, B, and C indicate the three extreme data points and correspond to the labeled rows in Table 8.1 Basic histograms and box plots allow you to rapidly identify extreme values in your data.

values beyond the upper or lower deciles (90th percentile, 10th percentile) of a distribution to be potential outliers, and those smaller than the 5th percentile or larger than the 95th percentile to be possible extreme values. Many statistical packages highlight these values on box plots (as we do with stars and letters in Figure 8.1) to indicate that you should scrutinize them carefully. Some of these packages also have interactive tools, described in the next section, that allow for location of these values within the spreadsheet. It is important to identify outliers because they can have dramatic effects on your statistical tests. Outliers and extreme values increase the variance in the data. Inflated variances decrease the power of the test and increase the chance of a Type II error (failure to reject a false null hypothesis; see Chapter 4). For this reason, some researchers automatically delete outliers and extreme values prior to conducting their analyses. This is bad practice! There are only two reasons for justifiably discarding data: (1) the data are in error (e.g., they were entered incorrectly in the field notebook); or (2) the data no longer represent valid observations from your original sample space (e.g., one of your dune plots was overrun by all-terrain vehicles). Deleting observations simply because they are “messy” is laundering or doctoring the results and legitimately could be considered scientific fraud. Outliers are more than just noise. Outliers can reflect real biological processes, and careful thought about their meaning can lead to new ideas and

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hypotheses.9 Moreover, some data points will appear as outliers only because the data are being forced into a normal distribution. Data from non-normal distributions such as the log-normal or exponential often seem to be extreme, but these data fall in place after they are appropriately transformed (see below). Errors

Errors are recorded values that do not represent the original measurements or observations. Some, but not all, errors are also outliers. Conversely, not all outliers in your data are necessarily errors. Errors can enter the dataset in two ways: as errors in collection or as errors in transcription. Errors in collection are data that result from broken or miscalibrated instruments, or from mistaken data entry in the field. Errors in collection can be difficult to identify. If you or your field assistant were recording the data and wrote down an incorrect value, the error rarely can be corrected.10 Unless the error is recognized immediately during data transcription, it will probably remain undetected and contribute to the variation of the data.

Although biologists are trained to think about means and averages, and to use statistics to test for patterns in the central tendency of data, interesting ecology and evolution often happen in the statistical tails of a distribution. Indeed, there is a whole body of statistics dedicated to the study and analysis of extreme values (Gaines and Denny 1993), which may have long-lasting impacts on ecosystems (e.g., Foster et al. 1998). For example, isolated populations (peripheral isolates, sensu Mayr 1963) that result from a single founder event may be sufficiently distinct in their genetic makeup that they would be considered statistical outliers relative to the original population. Such peripheral isolates, if subject to novel selection pressures, could become reproductively isolated from their parent population and form new species (Schluter 1996). Phylogenetic reconstruction has suggested that peripheral isolates can result in new species (e.g., Green et al. 2002), but experimental tests of the theory have not supported the model of speciation by founder effect and peripheral isolates (Mooers et al.1999). 10

To minimize collection errors in the field, we often engage in a call and response dialogue with each other and our field assistants: Data Collector: “This is Plant 107. It has 4 leaves, no phyllodes, no flowers.” Data Recorder: “Got it, 4 leaves no phyllodes or flowers for 107.” Data Collector: “Where’s 108?” Data Recorder: “ Died last year. 109 should be about 10 meters to your left. Last year it had 7 leaves and an aborted flower. After 109, there should be only one more plant in this plot.” Repeating the observations back to one another and keeping careful track of the replicates helps to minimize collection errors. This “data talk” also keeps us alert and focused during long field days.

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Errors resulting from broken or miscalibrated instruments are easier to detect (once it is determined that the instrument is broken or after it has been recalibrated). Unfortunately, instrument error can result in the loss of a large amount of data. Experiments that rely on automated data collection need to have additional procedures for checking and maintaining equipment to minimize data loss. It may be possible to collect replacement values for data resulting from equipment failures, but only if the failures are detected very soon after the data are collected. Potential equipment failures are another reason to transcribe rapidly your raw data into spreadsheets and evaluate their accuracy. Errors in transcription result primarily from mistyping values into spreadsheets. When these errors show up as outliers, they can be checked against the original field data sheets. Probably the most common transcription error is the misplaced decimal point, which changes the value by an order of magnitude. When errors in transcription do not show up as outliers, they can remain undetected unless spreadsheets are proofread very carefully. Errors in transcription are less common when data are transferred electronically from instruments directly into spreadsheets. However, transmission errors can occur and can result in “frame-shift” errors, in which one value placed in the wrong column results in all of the subsequent values being shifted into incorrect columns. Most instruments have built-in software to check for transmission errors and report them to the user in real-time. The original data files from automated data-collection instruments should not be discarded from the instrument’s memory until after the spreadsheets have been checked carefully for errors. Missing Data

A related, but equally important, issue is the treatment of missing data. Be very careful in your spreadsheets and in your metadata to distinguish between measured values of 0 and missing data (replicates for which no observations were recorded). Missing data in a spreadsheet should be given their own designation (such as the abbreviation “NA” for “not available”) rather than just left as blank cells. Be careful because some software packages may treat blank cells as zeroes (or insert 0s in the blanks), which will ruin your analyses. Detecting Outliers and Errors

We have found three techniques to be especially useful for detecting outliers and errors in datasets: calculating column statistics, checking ranges and precision of column values; and graphical exploratory data analysis. Other, more complex methods are discussed by Edwards (2000). We use as our example for outlier detection the measurement of heights of pitcher-leaves of Darlingtonia californica

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(the cobra lily, or California pitcher plant).11 We collected these data, reported in Table 8.1, as part of a study of the growth, allometry, and photosynthesis of this species (Ellison and Farnsworth 2005). In this example of only 25 observations, the outliers are easy to spot simply by scanning the columns of numbers. Most ecological datasets, however, will have far more observations and it will be harder to find the unusual values in the data file. COLUMN STATISTICS The calculation of simple column statistics within the spreadsheet is a straightforward way to identify unusually large or small values in the dataset. Measures of location and spread, such as the column mean, median, standard deviation, and variance, give a quick overview of the distribution of the values in the column. The minimum and maximum values of the column may indicate suspiciously large or small values. Most spreadsheet software packages have functions that calculate these values. If you enter these functions as the last six rows of the spreadsheet (Table 8.1), you can use them as a first check on the data. If you find an extreme value and determine that it is a true error, the spreadsheet will automatically update the calculations when you replace it with the correct number.

Another way to check the data is to use spreadsheet functions to ensure that values in a given column are within reasonable boundaries or reflect the precision of measurement. For example, Darlingtonia pitchers rarely exceed 900 mm in height. Any measurement much above this value would be suspect. We could look over the dataset carefully to see if any values are extremely large, but for real datasets with hundreds or thousands of observations, manual checking is inefficient and inaccurate. Most spreadsheets have logical functions that can be used for value checking and that can automate this process. Thus, you could use the statement: CHECKING RANGE AND PRECISION OF COLUMN VALUES

If (value to be checked) > (maximum value), record a “1”; otherwise, record a “0” to quickly check to see if any of the values of height exceed an upper (or lower) limit.

Darlingtonia californica

11 Darlingtonia californica is a carnivorous plant in the family Sarraceniaceae that grows only in serpentine fens of the Siskiyou Mountains of Oregon and California and the Sierra Nevada Mountains of California. Its pitcher-shaped leaves normally reach 800 mm in height, but occasionally exceed 1 m. These plants feed primarily on ants, flies, and wasps.

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TABLE 8.1 Ecological data set illustrating typical extreme values of Darlingtonia californica Plant # Height (mm) Mouth(mm) Tube (mm)

Mean

1 2 3 4 5 6 7 8 9 10

744 700 714 667 600 777 640 440 715 573

34.3 34.4 28.9 32.4 29.1 33.4 34.5 29.4 39.5 33.0

18.6 20.9 19.7 19.5 17.5 21.1 18.6 18.4 19.7 15.8

1500

33.8

19.1

12 13 14 15 16 17 18 19 20 21

650 480 545 845 560 450 600 607 675 550

36.3 27.0 30.3 37.3 42.1 31.2 34.6 33.5 31.4 29.4

20.2 18.1 17.3 19.3 14.6 20.6 17.1 14.8 16.3 17.6

0.3

0.1

30.2 35.8

16.5 15.7

5.1

23 24

534 655

65.5

611.7 Median 607 Standard deviation 265.1 Variance 70271 Minimum 5.1 Maximum 1500

3.52

The data are morphological measurements of individual pitcher plants. Pitcher height, pitcher mouth diameter, and pitcher tube diameter were recorded for 25 individuals sampled at Days Gulch. Rows with extreme values are shown in boldface and designated as A, B, and C, and are noted with asterisks in Figure 8.1. Summary statistics for each variable are given at the bottom of the table. It is crucial to identify and evaluate extreme values in a dataset. Such values greatly inflate variance estimates, and it is important to ascertain whether those observations represent measurement error, atypical measurements or simply natural variation of the sample. Data from Ellison and Farnsworth (2005).

1.77

30.6 33.0 9.3 86.8 0.3

16.8 18.1 5.1 26.1 0.1

42.1

21.1

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This same method can be used to check the precision of the values. We measured height using a measuring tape marked in millimeters, and routinely recorded measurements to the nearest whole millimeter. Thus, no value in the spreadsheet should be more precise than a single millimeter (i.e., no value should have fractions of millimeters entered). The logical statement to check this is: If (value to be checked has any value other than 0 after the decimal point), record a “1”; otherwise, record a “0” There are many similar checks on column values that you can imagine, and most can be automated using logical functions within spreadsheets. Range checking on the data in Table 8.1 reveals that the recorded height of Plant 11 (1500 mm) is suspiciously large because this observation exceeds the 900-mm threshold. Moreover the recorded heights of Plant 22 (5.1 mm) and Plant 25 (65.5 mm) were misentered because the decimal values are greater than the precision of our measuring tapes. This audit would lead us to re-examine our field data sheets or perhaps return to the field to measure the plants again. Scientific pictures—graphs—are one of the most powerful tools you can use to summarize the results of your study (Tufte 1986, 1990; Cleveland 1985, 1993). They also can be used to detect outliers and errors, and to illuminate unexpected patterns in your data. The use of graphics in advance of formal data analysis is called graphical exploratory data analysis or graphical EDA, or simply EDA (e.g., Tukey 1977; Ellison 2001). Here, we focus on the use of EDA for detecting outliers and extreme values. The use of EDA to hunt for new or unexpected patterns in the data (“data mining” or more pejoratively, “data dredging”) is evolving into its own cottage industry (Smith and Ebrahim 2002).12

GRAPHICAL EXPLORATORY DATA ANALYSIS

The statistical objection to data dredging is that if we create enough graphs and plots from a large, complex dataset, we will surely find some relationships that turn out to be statistically significant. Thus, data dredging undermines our calculation of P-values and increases the chances of incorrectly rejecting the null hypothesis (Type I error). The philosophical objection to EDA is that you should already know in advance how your data are going to be plotted and analyzed. As we emphasized in Chapters 6 and 7, laying out the design and analysis ahead of time goes hand-in-hand with having a focused research question. On the other hand, EDA is essential for detecting outliers and errors. And there is no denying that you can uncover interesting patterns just by casually plotting your data. You can always return to the field to test these patterns with an appropriate experimental design. Good statistical packages facilitate EDA because graphs can be constructed and viewed quickly and easily.

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0 1 2 3 4 H 5 M 6 H 7 8 9 10 11 12 13 14 15

458 34567 00145577 01148 4

219

Figure 8.2 Stem-and-leaf plot. The data are measurements of the heights (in mm) of 25 individuals of Darlingtonia californica (see Table 8.1). In this plot, the first column of numbers on the left—the “stem”—represent the “hundreds” place, and each “leaf” to its right represents the “tens” place of each recorded value. For example, the first row illustrates the observations 00x and 06x, or an observation less than 10 (the value 5.1 of Plant 22 of Table 8.1), and another observation between 60 and 70 (the value 65.5 of Plant 25). The fifth row illustrates observations 44x, 45x, and 48x, which correspond to values 440 (Plant 8), 450 (Plant 17), and 480 (Plant 13). The last row illustrates observation 150x, corresponding to value 1500 (Plant 11). All observations in column 1 of Table 8.1 are included in this display. The median (607) is in row 7, which is labeled with the letter M. The lower quartile (545), also known as a hinge, is in row 6, which is labeled with the letter H. The upper quartile or hinge (700) is in row 8, also labeled H. Like histograms and box plots, stem-and-leaf plots are a diagnostic tool for visualizing your data and detecting extreme data points.

There are three indispensable types of graphs for detecting outliers and extreme values: box plots, stem-and-leaf plots, and scatterplots. The first two are best used for univariate data (plotting the distribution of a single variable), whereas scatterplots are used for bivariate or multivariate data (plotting relationships among two or more variables). You have already seen many examples of univariate data in this book: the 50 observations of spider tibial spine length (see Figure 2.6 and Table 3.1), the presence or absence of Rhexia in Massachusetts towns (see Chapter 2), and the lifetime reproductive output of orange-spotted whirligig beetles (see Chapter 1) are all sets of measurements of a single variable. To illustrate outlier detection and univariate plots for EDA, we use the first column of data in Table 8.1—pitcher height. Our summary statistics in Table 8.1 suggest that there may be some problems with the data. The variance is large relative to the mean, and both the minimum and the maximum value look extreme. A box plot (top of Figure 8.1) shows that two points are unusually small and one point is unusually large. We can identify these unusual values in more detail in a stem-and-leaf plot (Figure 8.2), which is a variant of the more familiar histogram (see Chapter 1; Tukey 1977). The stem-and-leaf plot shows clearly that the two small values are an order of magnitude (i.e., approximately tenfold) smaller than the mean or median, and that the one large value is more than twice as large as the mean or median. How unusual are these values? Some Darlingtonia pitchers are known to exceed a meter in height, and very small pitchers can be found as well. Perhaps these three observations simply reflect the inherent variability in the size of cobra lily pitchers. Without additional information, there is no reason to think that these values are in error or are not part of our sample space, and it would be inappropriate to remove them from the dataset prior to further statistical analysis.

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We do have additional information, however. Table 8.1 shows two other measurements we took from these same pitchers: the diameter of the pitcher’s tube, and the diameter of the opening of the pitcher’s mouth. We would expect all of these variables to be related to each other, both because of previous research on Darlingtonia (Franck 1976) and because the growth of plant (and animal) parts tends to be correlated (Niklas 1994).13 A quick way explore how these variables co-vary with one another is to plot all possible pair-wise relationships among them (Figure 8.3). The scatterplot matrix shown in Figure 8.3 provides additional information with which to decide whether our unusual values are outliers. The two scatterplots in the top row of Figure 8.3 suggest that the unusually tall plant in Table 13

Imagine a graph in which you plotted the logarithm of pitcher length on the x-axis and the logarithm of tube diameter on the y-axis. This kind of allometric plot reveals the way that the shape of an organism may change with its size. Suppose that small cobra lilies were perfect scaled-down replicas of large cobra lilies. In this case, increases in tube diameter are synchronous with increases in pitcher length, and the slope of the line (β1) = 1 (see Chapter 9). The growth is isometric, and the small plants look like miniatures of the large plants. On the other hand, suppose that small pitchers had tube diameters that were relatively (but not absolutely) larger than the tube diameters of large plants. As the plants grow in size, tube diameters increase relatively slowly compared to pitcher lengths. This would be a case of negative allometry, and it would be reflected in a slope value β1 < 1. Finally, if there is positive allometry, the tube diameter increases faster with increasing pitcher length (β1 > 1). These patterns are illustrated by Franck (1976). A more familiar example is the human infant, in which the head starts out relatively large, but grows slowly, and exhibits negative allometry. Other body parts, such as fingers and toes, exhibit positive allometry and grow relatively rapidly. Allometric growth (either positive or negative) is the rule in nature. Very few organisms grow isometrically, with juveniles appearing as miniature replicas of adults. Organisms change shape as they grow, in part because of basic physiological constraints on metabolism and uptake of materials. Imagine that (as a simplification) the shape of an organism is a cube with a side of length L. The problem is that, as organisms increase in size, volume increases as a cubic function (L3) of length (L), but surface area increases only as a square function (L2). Metabolic demands of an organism are proportional to volume (L3), but the transfer of material (oxygen, nutrients, waste products) is proportional to surface area (L2). Therefore, structures that function well for material transfer in small organisms will not get the job done in larger organisms. This is especially obvious when we compare species of very different body size. For example, the cell membrane of a tiny microorganism does a fine job transferring oxygen by simple diffusion, but a mouse or a human requires a vascularized lung, circulatory system, and a specialized transport molecule (hemoglobin) to accomplish the same thing. Both within and between species, patterns of shape, size, and morphology often reflect the necessity of boosting surface area to meet the physiological demands of increasing volume. See Gould (1977) for an extended discussion of the evolutionary significance of allometric growth.

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40 A

1500 1000

Height 500 BC

40 A A

Mouth

20 10 C B

Tube diameter (mm)

Mouth diameter (mm)

Pitcher height (mm)

C 20

15 Tube

10 5

C B 0

B 500 1000 1500 Pitcher height (mm)

0 0

Mouth diameter (mm)

20 5 10 15 Tube diameter (mm)

Figure 8.3 Scatterplot matrix illustrating the relationship between the height, mouth diameter, and tube diameter of 25 D. californica individuals (see Table 8.1). The extreme values labeled in Figure 8.1 and Table 8.1 are also labeled here. The diagonal panels in this figure illustrate the histograms for each of the 3 morphological variables. In the off-diagonal panels, scatterplots illustrate the relationship between the two variables indicated in the diagonal; the scatterplots above the diagonal are mirror images of those below the diagonal. For example, the scatterplot at the lower left illustrates the relationship between pitcher height (on the x-axis) and tube diameter (on the y-axis), whereas the scatterplot at the upper right illustrates the relationship between tube diameter (on the x-axis) and pitcher height (on the y-axis). The lines in each scatterplot are the best-fit linear regressions between the two variables (see Chapter 9 for more discussion of linear regression).

8.1 (Point A; Plant 11) is remarkably tall for its tube and mouth diameters. The plots of mouth diameter versus tube diameter, however, show that neither of these measurements are unusual for Plant 11. Collectively, these results suggest an error either in recording the plant’s height in the field or in transcribing its height from the field datasheet to the computer spreadsheet.

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In contrast, the short plants are not unusual in any regard except for being short. Not only are they short, but they also have small tubes and small mouths. If we remove Plant 11 (the unusually tall plant) from the dataset (Figure 8.4), the relationships among all three variables of the short plants are not unusual relative to the rest of the population, and there is no reason based on the data themselves to suspect that the values for these plants were entered incorrectly. Nonetheless, these two plants are very small (could we really have measured a

Pitcher height (mm)

800 600 Height

400 200 B

40 30 Mouth

20 10 C

C 20

Tube diameter (mm)

Mouth diameter (mm)

15 Tube

10 5

C B 0

B 200 400 600 800 Pitcher height (mm)

0 0

Mouth diameter (mm)

5 10 15 20 Tube diameter (mm)

Figure 8.4 Scatterplot matrix illustrating the relationship between pitcher height, mouth diameter, and tube diameter. The data and figure layout are identical to Figure 8.3. However, the outlier Point A (see Table 8.1) has been eliminated, which changes some of these relationships. Because Point A was a large outlier for pitcher height (height = 1500 mm), the scale of this variable (x-axis in the left-hand column) now ranges from only 0 to 800 mm, compared to 0 to 1500 mm in Figure 8.3. It is not uncommon to see correlations and statistical significance change based on the inclusion or exclusion of a single extreme data value.

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5mm-tall plant?) and we should re-check our datasheets and perhaps return to the field to re-measure these plants. Creating an Audit Trail

The process of examining a dataset for outliers and errors is a special case of the generic quality assurance and quality control process (abbreviated QA/QC). As with all other operations on a spreadsheet or dataset, the methods used for outlier and error detection should be documented in a QA/QC section of the metadata. If it is necessary to remove data values from the spreadsheet, a description of the values removed should be included in the metadata, and a new spreadsheet should be created that contains the corrected data. This spreadsheet also should be stored on permanent media and given a unique identifier in the data catalog. The process of subjecting data to QA/QC and modifying data files leads to the creation of an audit trail. An audit trail is a series of files and their associated documentation that enables other users to recreate the process by which the final, analyzed dataset was created. The audit trail is a necessary part of the metadata for any study. If you are preparing an environmental impact report or other document that is used in legal proceedings, the audit trail may be part of the legal evidence supporting the case.14

The Final Step: Transforming the Data You will often read in a scientific paper that the data were “transformed prior to analysis.” Miraculously, after the transformation, the data “met the assumptions” of the statistical test being used and the analysis proceeded apace. Little else is usually said about the data transformations, and you may wonder why the data were transformed at all. But first, what is a transformation? By a transformation, we simply mean a mathematical function that is applied to all of the observations of a given variable: Y* = f(Y) 14

(8.1)

Historians of science have reconstructed the development of scientific theories by careful study of inadvertent audit trails—marginal notes scrawled on successive drafts of handwritten manuscripts and different editions of monographs. The use of word processors and spreadsheets has all but eliminated marginalia and yellow pads, but does allow for the creation and maintenance of formal audit trails. Other users of your data and future historians of science will thank you for maintaining an audit trail of your data.

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Y represents the original variable, Y* is the transformed variable, and f is a mathematical function that is applied to the data. Most transformations are fairly simple algebraic functions, subject to the requirement that they are continuous monotonic functions.15 Because they are monotonic, transformations do not change the rank order of the data, but they do change the relative spacing of the data points, and therefore affect the variance and shape of the probability distribution (see Chapter 2). There are two legitimate reasons to transform your data before analysis. First, transformations can be useful (although not strictly necessary) because the patterns in the transformed data may be easier to understand and communicate than patterns in the raw data. Second, transformations may be necessary so that the analysis is valid—this is the “meeting the assumptions” reason that is used most frequently in scientific papers and discussed in biometry textbooks. Data Transformations as a Cognitive Tool

Transformations often are useful for converting curves into straight lines. Linear relationships are easier to understand conceptually, and often have better statistical properties (see Chapter 9). When two variables are related to each other by multiplicative or exponential functions, the logarithmic transformation is one of the most useful data transformations (see Footnote 13). A classic ecological example is the species–area relationship: the relationship between species number and island or sample area (Preston 1962; MacArthur and Wilson 1967). If we measure the number of species on an island and plot it against the area of the island, the data often follow a simple power function: S = cAz

(8.2)

where S is the number of species, A is island area, and c and z are constants that are fitted to the data. For example, the number of species of plants recorded from each of the Galápagos Islands (Preston 1962) seems to follow a power relationship (Table 8.2). First, note that island area ranges over three orders of magnitude, from less than 1 to nearly 7500 km2. Similarly, species richness spans two orders of magnitude, from 7 to 325.

A continuous function is a function f(X) such that for any two values of the random variable X, xi and xj , that differ by a very small number (|xi – xj| < δ), then |f(xi) – f(xj)| < ε, another very small number. A monotonic function is a function f(X) such that for any two values of the random variable X, xi and xj , if xi < xj then f(xi)< f(xj ). A continuous monotonic function has both of these properties.

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TABLE 8.2 Species richness on 17 of the Galápagos Islands Island

Area (km2)

Number of species

log10 (Area)

log10 (Species)

Albemarle (Isabela) Charles (Floreana) Chatham (San Cristóbal) James (Santiago) Indefatigable (Santa Cruz) Abingdon (Pinta) Duncan (Pinzón) Narborough (Fernandina) Hood (Española) Seymour Barringon (Santa Fé) Gardner Bindloe (Marchena) Jervis (Rábida) Tower (Genovesa) Wenman (Wolf) Culpepper (Darwin)

5824.9 165.8 505.1 525.8 1007.5 51.8 18.4 634.6 46.6 2.6 19.4 0.5 116.6 4.8 11.4 4.7 2.3

325 319 306 224 193 119 103 80 79 52 48 48 47 42 22 14 7

3.765 2.219 2.703 2.721 3.003 1.714 1.265 2.802 1.669 0.413 1.288 –0.286 2.066 0.685 1.057 0.669 0.368

2.512 2.504 2.486 2.350 2.286 2.076 2.013 1.903 1.898 1.716 1.681 1.681 1.672 1.623 1.342 1.146 0.845

Mean Standard deviation Standard error of the mean

526.0 1396.9 338.8

119.3 110.7 26.8

1.654 1.113 0.270

1.867 0.481 0.012

Area (originally given in square miles) has been converted to square kilometers, and island names from Preston (1962) are retained, with modern island names given in parentheses. The last two columns give the logarithm of area and species richness, respectively. Mathematical transformations such as logarithms are used to better meet the assumptions of parametric analyses (normality, linearity, and constant variances) and they are used as a diagnostic tool to aid in the identification of outliers and extreme data points. Because both the response variable (species richness) and the predictor variable (island area) are measured on a continuous scale, a regression model is used for analysis (see Table 7.1).

If we plot the raw data—the number of species S as a function of area A (Figure 8.5)—we see that most of the data points are clustered to the left of the figure (as most islands are small). As a first pass at the analysis, we could try fitting a straight line to this relationship: S = β0 + β1A

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400

Number of species

226

300

200

100

0 0

1000 2000 3000 4000 5000 6000 Island area (km2)

Figure 8.5 Plot of plant species richness as a function of Galápagos Island area using the raw data (Columns 2 and 3) of Table 8.2. The line shows the best-fit linear regression line (see Chapter 9). Although a linear regression can be fit to any pair of continuous variables, the linear fit to these data is not good: there are too many negative outliers at small island areas, and the slope of the line is dominated by Albermarle, the largest island in the dataset. In many cases, a mathematical transformation of the X variable, the Y variable, or both will improve the fit of a linear regression.

In this case, β0 represents the intercept of the line, and β1 is its slope (see Chapter 9). However, the line does not fit the data very well. In particular, notice that the slope of the regression line seems to be dominated by the datum for Albermarle, the largest island in the dataset. In Chapter 7, we warned of precisely this problem that can arise with outlier data points that dominate the fit of a regression line (see Figure 7.3). The line fit to the data does not capture well the relationship between species richness and island area. If species richness and island area are related exponentially (see Equation 8.2), we can transform this equation by taking logarithms of both sides: log(S) = log(cAz)

(8.4)

log(S) = log(c) + zlog(A)

(8.5)

This transformation takes advantage of two properties of logarithms. First, the logarithm of a product of two numbers equals the sum of their logarithms: log(ab) = log(a) + log(b)

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(8.6)

The Final Step: Transforming the Data

Second, the logarithm of a number raised to a power equals the power times the logarithm of the number: log(ab) = blog(a)

(8.7)

We can rewrite Equation 8.4, denoting logarithmically transformed values with an asterisk (*): S* = c* + zA*

(8.8)

Thus, we have taken an exponential equation, Equation 8.2, and transformed it into a linear equation, Equation 8.8. When we plot the logarithms of the data, the relationship between species richness and island area is now much clearer (Figure 8.6), and the coefficients have a simple interpretation. The value for z in Equation 8.2 and 8.8, which equals the slope of the line in Figure 8.6, is 0.331; this means that every time we increase A* (the logarithm of island area) by one unit (that is, by a factor of 10 as we used log10 to transform our variables in Table 8.2, and 101 = 10), we increase species richness by 0.331 units (that is, by a factor of approximately 2 because 100.331 equals 2.14). Thus, we can say that a

log10(Number of species)

2.5

2.0

1.5

1.0

0.5 –1.0

0.0

1.0 2.0 log10(Island area)

3.0

4.0

Figure 8.6 Plot of the logarithm of plant species richness as a function of the logarithm of Galápagos Island area (Columns 4 and 5 of Table 8.2). The line is the best-fit linear regression line (see Chapter 9). Compared to the linear plot in Figure 8.5, this regression fits the data considerably better: the largest island in the data set no longer looks like an outlier, and the linearity of the fit is improved.

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10-fold increase in island area results in a doubling of the number of species present on the Galápagos Islands.16 Other transformations can be used to convert non-linear relationships to 3 linear ones. For example, the cube-root transformation ( 冑苳 Y ) is appropriate 3 for measures of mass or volume (Y ) that are allometrically related to linear measures of body size or length (Y; see Footnote 13). In studies that examine relationships between two measures of masses or volumes (Y 3), such as comparisons of brain mass and body mass, both the X and Y variables are logarithmically transformed. The logarithmic transformation reduces variation in data that may range over several orders of magnitude. (For a detailed exposition of the brain-body mass relationship, see Allison and Cicchetti 1976 and Edwards 2000.)

16 The history of the species-area relationship illustrates the dangers of reification: the conversion of an abstract concept into a material thing. The power function (S = cAz) has formed the basis for several important theoretical models of the speciesarea relationship (Preston 1962; MacArthur and Wilson 1967; Harte et al. 1999). It also has been used to argue for making nature reserves as large as possible so that they contain the greatest possible number of species. This led to a long debate on whether a single large or several small preserves would protect the most species (e.g., Willis 1984; Simberloff and Abele 1984). However, Lomolino and Weiser (2001) have recently proposed that the species-area relationship has an asymptote, in which case a power function is not appropriate. But this proposal has itself been challenged on both theoretical and empirical grounds (Williamson et al. 2001). In other words, there is no consensus that the power function always forms the basis for the speciesarea relationship (Tjørve 2003, 2009; Martin and Goldenfeld 2006). The power function has been popular because it appears to provide a good fit to many species-area datasets. However, a detailed statistical analysis of 100 published species-area relationships (Connor and McCoy 1979) found that the power function was the best-fitting model in only half of the datasets. Although island area is usually the single strongest predictor of species number, area typically accounts for only half of the variation in species richness (Boecklen and Gotelli 1984). As a consequence, the value of the species-area relationship for conservation planning is limited because there will be much uncertainty associated with species richness forecasts. Moreover, the species-area relationship can be used to predict only the number of species present, whereas most conservation management strategies will be concerned with the identity of the resident species. The moral of the story is that data can always be fit to a mathematical function, but we have to use statistical tools to evaluate whether the fit is reasonable or not. Even if the fit of the data is acceptable, this result, by itself, is rarely a strong test of a scientific hypothesis, because there are usually alternative mathematical models that can fit the data just as well.

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The Final Step: Transforming the Data

Data Transformations because the Statistics Demand It

All statistical tests require that the data fit certain mathematical assumptions. For example, data to be analyzed using analysis of variance (see Chapters 7 and 10) must meet two assumptions: 1. The data must be hom*oscedastic—that is, the residual variances of all treatment groups need to be approximately equal to each other. 2. The residuals, or deviations from the means of each group, must be normal random variables. Similarly, data to be analyzed using regression or correlation (see Chapter 9) also should have normally distributed residuals that are uncorrelated with the independent variable. Mathematical transformations of the data can be used to meet these assumptions. It turns out that common data transformations often address both assumptions simultaneously. In other words, a transformation that equalizes variances (Assumption 1) often normalizes the residuals (Assumption 2). Five transformations are used commonly with ecological and environmental data: the logarithmic transformation, the square-root transformation, the angular (or arcsine) transformation, the reciprocal transformation, and the BoxCox transformation. The logarithmic transformation (or log transformation) replaces the value of each observation with its logarithm:17

THE LOGARITHMIC TRANSFORMATION

Logarithms were invented by the Scotsman John Napier (1550–1617). In his epic tome Mirifici logarithmorum canonis descriptio (1614), he sets forth the rationale for using logarithms (from the English translation, 1616): Seeing there is nothing (right well-beloved Students of the Mathematics) that is so troublesome to mathematical practice, nor that doth more molest and hinder calculators, than the multiplications, divisions, square and cubical extractions of great numbers, which John Napier besides the tedious expense of time are for the most part subject to many slippery errors, I began therefore to consider in my mind by what certain and ready art I might remove those hindrances. As illustrated in Equations 8.4–8.8, the use of logarithms allows multiplication and division to be carried out by simple addition and subtraction. Series of numbers that are multiplicative on a linear scale turn out to be additive on a logarithmic scale (see Footnote 2 in Chapter 3).

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Y* = log(Y)

(8.9)

The logarithmic transformation (most commonly using the natural logarithm or the logarithm to the base e)18 often equalizes variances for data in which the mean and the variance are positively correlated. A positive correlation means that groups with large averages will also have large variances (in ANOVA) or that the magnitude of the residuals is correlated with the magnitude of the independent variable (in regression). Univariate data that are positively skewed (skewed to the right) often have a few large outlier values. With a log-transformation, these outliers often are drawn into the mainstream of the distribution, which becomes more symmetrical. Datasets in which means and variances are positively correlated also tend to have outliers with positively skewed residuals. A logarithmic transformation often addresses both problems simultaneously. Note that the logarithm of 0 is not defined: regardless of base b, there is no number for which ab = 0. One way around this problem is to add 1 to each observation before taking its logarithm (as log(1) = 0, again regardless of base b). However, this is not a useful or appropriate solution if the dataset (or especially if one treatment group) contains many zeros.19 THE SQUARE-ROOT TRANSFORMATION The square-root transformation replaces the value of each observation with its square root:

Y* = 冑苳苳 Y

(8.10)

18 Logarithms can be taken with respect to many “bases,” and Napier did not specify a particular base in his Mirifici. In general, for a value a and a base b we can write

logba = X X

which means that b = a. We have already used the logarithm “base 10” in our example of the species-area relationship in Table 8.2; for the first line in Table 8.2, the log10(5825) = 3.765 because 103.765 = 5825 (allowing for round-off error). A change in one log10 unit is a power of 10, or an order of magnitude. Other logarithmic bases used in the ecological literature are 2 (the octaves of Preston (1962) and used for data that increase by powers of 2), and e, the base of the natural logarithm, or approximately 2.71828… e is a transcendental number, whose value was proven by Leonhard Euler (1707–1783) to equal 1 lim(1 + )n n

n→∞

The loge was first referred to as a “natural logarithm” by the mathematician Nicolaus Mercator (1620–1687), who should not to be confused with the map-maker Gerardus Mercator (1512–1594). It is often written as ln instead of loge.

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The Final Step: Transforming the Data

This transformation is used most frequently with count data, such as the number of caterpillars per milkweed or the number of Rhexia per town. In Chapter 2 we showed that such data often follow a Poisson distribution, and we pointed out that the mean and variance of a Poisson random variable are equal (both equal the Poisson rate parameter λ). Thus, for a Poisson random variable, the mean and variance vary identically. Taking square-roots of Poisson random variables yields a variance that is independent of the mean. Because the square root of 0 itself equals 0, the square-root transformation does not transform data values that are equal to 0. Thus, to complete the transformation, you should add some small number to the value prior to taking its square root. Adding 1/2 (0.5) to each value is suggested by Sokal and Rohlf (1995), whereas 3/8 (0.325) is suggested by Anscombe (1948). THE ARCSINE OR ARCSINE-SQUARE ROOT TRANSFORMATION The arcsine, arcsinesquare root, or angular transformation replaces the value of each observation with the arcsine of the square root of the value:

Y* = arcsine 冑苳苳 Y

(8.11)

This transformation is used principally for proportions (and percentages), which are distributed as binomial random variables. In Chapter 3, we noted that the mean of a binomial distribution = np and its variance = np(1 – p), where p is the probability of success and n is the number of trials. Thus, the variance is a direct function of the mean (the variance = (1 – p) times the mean). The arcsine transformation (which is simply the inverse of the sine function) removes this dependence. Because the sine function yields values only between –1 and +1, the inverse sine function can be applied only to data whose values fall 19

Adding a constant before taking logarithms can cause problems for estimating population variability (McArdle et al. 1990), but it is not a serious issue for most parametric statistics. However, there are some subtle philosophical issues in how zeroes in the data are treated. By adding a constant to the data, you are implying that 0 represents a measurement error. Presumably, the true value is some very small number, but, by chance, you measured 0 for that replicate. However, if the value is a true zero, the most important source of variation may be between the absence and presence of the quantity measured. In this case, it would be more appropriate to use a discrete variable with two categories to describe the process. The more zeros in the dataset (and the more that are concentrated in one treatment group), the more likely it is that 0 represents a qualitatively different condition than a small positive measurement. In a population time series, a true zero would represent a population extinction rather than just an inaccurate measurement.

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between –1 and +1: –1 ≤ Yi ≤ +1. Therefore, this transformation is appropriate only for data that are expressed as proportions (such as p, the proportion or probability of success in a binomial trial). Two caveats to note with the arcsine transformation. First, if your data are percentages (0 to 100 scale), they must first be converted to proportions (0 to 1.0 scale). Second, in most software packages, the arcsine function gives transformed data in units of radians, not degrees. THE RECIPROCAL TRANSFORMATION The reciprocal transformation replaces the value of each observation with its reciprocal:

Y* = 1/Y

(8.12)

It is used most commonly for data that record rates, such as number of offspring per female. Rate data often appear hyperbolic when plotted as a function of the variable in the denominator. For example if you plot the number of offspring per female on the y-axis and number of females in the population on the x-axis, the resulting curve may resemble a hyperbola, which decreases steeply at first, then more gradually as X increases. These data are generally of the form aXY = 1 (where X is number of females and Y is number of offspring per female), which can be re-written as a hyperbola 1/Y = aX Transforming Y to its reciprocal 1/Y results in a new relationship Y* = 1/(1/Y) = aX which is more amenable to linear regression. THE BOX-COX TRANSFORMATION

We use the logarithmic, square root, reciprocal and other transformations to reduce variance and skew in the data and to create a transformed data series that has an approximately normal distribution. The final transformation is the Box-Cox transformation, or generalized power transformation. This transformation is actually a family of transformations, expressed by the equation Y* = (Yλ – 1)/λ Y* = loge(Y)

(for λ ≠ 0) (for λ = 0)

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(8.13)

The Final Step: Transforming the Data

where λ is the number that maximizes the log-likelihood function: ν ν n L = − log e (sT2 ) + (λ − 1) ∑ log eY n i=1 2

(8.14)

where ν is the degrees of freedom, n is the sample size, and sT2 is the variance of the transformed values of Y (Box and Cox 1964). The value of λ that results when Equation 8.14 is maximized is used in Equation 8.13 to provide the closest fit of the transformed data to a normal distribution. Equation 8.14 must be solved iteratively (trying different values of λ until L is maximized) with computer software. Certain values of λ correspond to the transformations we have already described. When λ = 1, Equation 8.13 results in a linear transformation (a shift operation; see Figure 2.7), when λ = 1/2, the result is the square-root transformation, when λ = 0, the result is the natural logarithmic transformation, and when λ = –1, the result is the reciprocal transformation. Before going to the trouble of maximizing Equation 8.14, you should try transforming your data using simple arithmetic transformations. If your data are right-skewed, try using the more familiar transformations from the series 1/冑苳苳 Y , 冑苳苳 Y , ln(Y), 1/Y. If your data 2 3 are left-skewed, try Y , Y , etc. (Sokal and Rohlf 1995). Reporting Results: Transformed or Not?

Although you may transform the data for analysis, you should report the results in the original units. For example, the species–area data in Table 8.2 would be analyzed using the log-transformed values, but in describing island size or species richness, you would report them in their original units, back-transformed. Thus, the average island size is –––– antilog(log A ) = antilog(1.654) = 45.110 km2 Note that this is very different from the arithmetic average of island size prior to transformation, which equals 526 km2. Similarly, average species richness is –––– antilog(log S ) = 101.867 = 73.6 species which differs from the arithmetic average of species richness prior to transformation, 119.3. You would construct confidence intervals (see Chapter 3) in a similar manner, by taking antilogarithms of the confidence limits constructed using the standard errors of the means of the transformed data. This will normally result in asymmetric confidence intervals. For the island area data in Table 8.2, the stan-

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Raw data (field notebooks, data loggers, instruments, etc.)

Detailed methods (field notebooks, data collection software, etc.)

Organize data into spreadsheets

Generate initial (basic) metadata Documented (and useable) dataset

Store archival version of dataset on permanent media (paper and other optical media, e.g., CD-ROM)

Dataset checked for outliers and errors?

YES Document transformation; add to metadata; update audit trail

Transform data

Exploratory data analysis Summary statistics

YES

Do data require transformation? NO

Store archival version of dataset to be analyzed on permanent media

Proceed to data analysis

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Document QA/QC procedures, add to metadata with date stamp (create audit trail)

QA/QC check and flag outliers. Are outliers valid data or the result of errors in collection or transcription?

Summary: The Data Management Flow Chart

Figure 8.7 The data management flowchart. This chart outlines the steps for managing, curating, and storing data. These steps should all be taken before formal data analysis begins. Documentation of metadata is especially important to ensure that the dataset will still be accessible and usable in the distant future.

dard error of the mean of log(A) = 0.270. For n = 17, the necessary value from the t distribution, t0.025[16] = 2.119. Thus, the lower bound of the 95% confidence interval (using Equation 3.16) = 1.654 – 2.119 × 0.270 = 1.082. Similarly, the upper bound of the 95% confidence interval = 1.654 + 2.119 × 0.270 = 2.226. On a logarithmic scale these two values form a symmetrical interval around the mean of 1.654, but when they are back-transformed, the interval is no longer symmetrical. The antilog of 1.082 = 101.082 = 12.08, whereas the antilog of 2.226 = 102.226 = 168.27; the interval [12.08, 168.27] which is not symmetrical around the back-transformed mean of 45.11. The Audit Trail Redux

The process of transforming data also should be added to your audit trail. Just as with data QA/QC, the methods used for data transformation should be documented in the metadata. Rather than writing over your data in your original spreadsheet with the transformed values, you should create a new spreadsheet that contains the transformed data. This spreadsheet also should be stored on permanent media, given a unique identifier in your data catalog, and added to the audit trail.

Summary: The Data Management Flow Chart Organization, management, and curation of your dataset are essential parts of the scientific process, and must be done before you begin to analyze your data. This overall process is summarized in our data management flow chart (Figure 8.7). Well-organized datasets are a lasting contribution to the broader scientific community, and their widespread sharing enhances collegiality and increases the pace of scientific progress. The free and public availability of data is a requirement of many granting agencies and scientific journals. Data collected with public funds distributed by organizations and agencies in the United States may be requested at any time by anyone under the provisions of the Freedom of Information Act. Adequate funds should be budgeted to allow for necessary and sufficient data management. Data should be organized and computerized rapidly following collection, documented with sufficient metadata necessary to reconstruct the data collection, and archived on permanent media. Datasets should be checked carefully

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for errors in collection and transcription, and for outliers that are nonetheless valid data. Any changes or modifications to the original (“raw”) data resulting from error- and outlier-detection procedures should be documented thoroughly in the metadata; the altered files should not replace the raw data file. Rather, they should be stored as new (modified) data files. An audit trail should be used to track subsequent versions of datasets and their associated metadata. Graphical exploratory data analysis and calculation of basic summary statistics (e.g., measures of location and spread) should precede formal statistical analysis and hypothesis testing. A careful examination of preliminary plots and tables can indicate whether or not the data need to be transformed prior to further analysis. Data transformations also should be documented in the metadata. Once a final version of the dataset is ready for analysis, it too should be archived on permanent media along with its complete metadata and audit trail.

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PART III

Data Analysis

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CHAPTER 9

Regression

Regression is used to analyze relationships between continuous variables. At its most basic, regression describes the linear relationship between a predictor variable, plotted on the x-axis, and a response variable, plotted on the y-axis. In this chapter, we explain how the method of least-squares is used to fit a regression line to data, and how to test hypotheses about the parameters of the fitted model. We highlight the assumptions of the regression model, describe diagnostic tests to evaluate the fit of the data to the model, and explain how to use the model to make predictions. We also provide a brief description of some more advanced topics: logistic regression, multiple regression, non-linear regression, robust regression, quantile regression, and path analysis. We close with a discussion of the problems of model selection: how to choose an appropriate subset of predictor variables, and how to compare the relative fit of different models to the same data set.

Defining the Straight Line and Its Two Parameters We will start with the development of a linear model, because this is the heart of regression analysis. As we noted in Chapter 6, a regression model begins with a stated hypothesis about cause and effect: the value of the X variable causes, either directly or indirectly, the value of the Y variable.1 In some cases, the direc1

Many statistics texts emphasize a distinction between correlation, in which two variables are merely associated with one another, and regression, in which there is a direct cause-and-effect relationship. Although different kinds of statistics have been developed for both cases, we think the distinction is largely arbitrary and often just semantic. After all, investigators do not seek out correlations between variables unless they believe or suspect there is some underlying cause-and-effect relationship. In this chapter, we cover statistical methods that are sometimes treated separately in analyses of correlation and regression. All of these techniques can be used to estimate and test associations of two or more continuous variables.

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tion of cause and effect is straightforward—we hypothesize that island area controls plant species number (see Chapter 8) and not the other way around. In other cases, the direction of cause and effect is not so obvious—do predators control the abundance of their prey, or does prey abundance dictate the number of predators (see Chapter 4)? Once you have made the decision about the direction of cause and effect, the next step is to describe the relationship as a mathematical function: Y = f(X)

(9.1)

In other words, we apply the function f to each value of the variable X (the input) to generate the corresponding value of the variable Y (the output). There are many interesting and complex functions that can describe the relationship between two variables, but the simplest one is that Y is a linear function of X: Y = β0 + β1X

(9.2)

In words, this function says “take the value of the variable X, multiply it by β1, and add this number to β0. The result is the value of the variable Y.” This equation describes the graph of a straight line. This model has two parameters in it, β0 and β1, which are called the intercept and the slope of the line (Figure 9.1). The intercept (β0) is the value of the function when X = 0. The intercept is measured in the same units as the Y variable. The slope (β1) measures the change in the Y variable for each unit change in the X variable. The slope is therefore a rate and is measured in units of ΔY/ΔX (read as: “change in Y divided by change in X”). If the slope and intercept are known, Equation 9.2 can be used to predict the value of Y for any value of X. Conversely, Equation 9.2

β1 Y

Figure 9.1 Linear relationship between variables X and Y. The line is described by the equation y = β0 + β1X, where β0 is the intercept, and β1 is the slope of the line. The intercept β0 is the predicted value from the regression equation when X = 0. The slope of the line β1 is the increase in the Y variable associated with a unit increase in the X variable (ΔY/ΔX). If the value of X is known, the predicted value of Y can be calculated by multiplying X by the slope (β1) and adding the intercept (β0).

1.0

β0

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Fitting Data to a Linear Equation

Extrapolation

Interpolation

Extrapolation

Figure 9.2 Linear models may approximate non-linear functions over a limited domain of the X variable. Interpolation within these limits may be acceptably accurate, even though the linear model (blue line) does not describe the true functional relationship between Y and X (black curve). Extrapolations will become increasingly inaccurate as the forecasts move further away from the range of collected data. A very important assumption of linear regression is that the relationship between X and Y (or transformations of these variables) is linear.

can be used to determine the value of X that would have generated a particular value of Y. Of course, nothing says that nature has to obey a linear equation; many ecological relationships are inherently non-linear. However, the linear model is the simplest starting place for fitting functions to data. Moreover, even complex, non-linear (wavy) functions may be approximately linear over a limited range of the X variable (Figure 9.2). If we carefully restrict our conclusions to that range of the X variable, a linear model may be a valid approximation of the function.

Fitting Data to a Linear Model Let’s first recast Equation 9.2 in a form that matches the data we have collected. The data for a regression analysis consist of a series of paired observations. Each observation includes an X value (Xi) and a corresponding Y value (Yi) that both have been measured for the same replicate. The subscript i indicates the replicate. If there is a total of n replicates in our data, the subscript i can take on any integer value from i = 1 to n. The model we will fit is Yi = β 0 + β1 X i + ε i

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(9.3)

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As in Equation 9.2, the two parameters in the linear equation, β0 and β1, are unknowns. But there is also now a third unknown quantity, εi, which represents the error term. Whereas β0 and β1 are simple constants, εi is a normal random variable. This distribution has an expected value (or mean) of 0, and a variance equal to σ2, which may be known or unknown. If all of the data points fall on a perfectly straight line, then σ2 equals 0, and it would be an easy task to connect those points and then measure the intercept (β0) and the slope (β1) directly from the line.2 However, most ecological datasets exhibit more variation than this—a single variable rarely will account for most of the variation in the data, and the data points will fall within a fuzzy band rather than along a sharp line. The larger the value of σ2, the more noise, or error, there will be about the regression line. In Chapter 8, we illustrated a linear regression for the relationship between island area (the X variable) and plant species number (the Y variable) in the Galápagos Islands (see Figure 8.6). Each point in Figure 8.6 consisted of a paired observation of the area of the island and its corresponding number of plant species (see Table 8.2). As we explained in Chapter 8, we used a logarithmic transformation of both the X variable (area) and the Y variable (species richness) in order to hom*ogenize the variances and linearize the curve. A little later in this chapter, we will return to these data in their untransformed state. Figure 8.6 showed a clear relationship between log10(area) and log10(species number), but the points did not fall in a perfect line. Where should the regression line be placed? Intuitively, it seems that the regression line should pass – – through the center of the cloud of data, defined by the point (X , Y ). For the island data, the center corresponds to the point (1.654, 1.867) (remember, these are log-transformed values). Now we can rotate the line through that center point until we arrive at the “best fit” position. But how should we define the best fit for the line? Let’s first define the squared residual di2 as the squared difference between the observed Yi value and the Y value that is predicted by the regression equation (Yˆi). We use the small caret (or “hat”) to distinguish between the observed value (Yi) and the value predicted from the regression equation (Yˆi). The squared residual di2 is calculated as di2 = (Yi − Yˆi )2 2

(9.4)

Of course, if there are only two observations, a straight line will fit them perfectly every time! But having more data is no guarantee that the straight line is a meaningful model. As we will see in this chapter, a straight line can be fit to any set of data and used to make forecasts, regardless of whether the fitted model is valid or not. However, with a large data set, we have diagnostic tools for assessing the fit of the data, and we can assign confidence intervals to forecasts that are derived from the model.

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Fitting Data to a Linear Equation

The residual deviation is squared because we are interested in the magnitude, and not the sign, of the difference between the observed and predicted value (see Footnote 8 in Chapter 2). For any particular observation Yi , we could pass the regression through that point, so that its residual would be minimized (di = 0). But the regression line has to fit all of the data collectively, so we will define the sum of all of the residuals, also called the residual sum of squares and abbreviated as RSS, to be n

RSS = ∑ (Yi − Yˆi )2

(9.5)

i=1

The “best fit” regression line is the one that minimizes the residual sum of squares.3 By minimizing RSS, we are ensuring that the regression line results in the smallest average difference between each Yi value and the Yˆi value that is predicted by the regression model (Figure 9.3).4 3

Francis Galton (1822–1911)—British explorer, anthropologist, cousin of Charles Darwin, sometime statistician, and quintessential D.W.E.M. (Dead White European Male)—is best remembered for his interest in eugenics and his assertion that intelligence is inherited and little influenced by environmental factors. His writings on intelligence, race, and heredity led him to advocate breeding restrictions among people and undergirded early racist policies in colonial Australia. He was knighted in 1909. Francis Galton In his 1866 article “Regression towards mediocrity in hereditary structure,” Galton analyzed the heights of adult children and their parents with the least-squares linear regression model. Although Galton’s data frequently have been used to illustrate linear regression and regression toward the mean, a recent re-analysis of the original data reveals they are not linear (Wachsmuth et al. 2003). Galton also invented a device called the quincunx, a box with a glass face, rows of pins inside, and a funnel at the top. Lead shot of uniform size and mass dropped through the funnel was deflected by the pins into a modest number of compartments. Drop enough shot, and you get a normal (Gaussian) curve. Galton used this device to obtain empirical data showing that the normal curve can be expressed as a mixture of other normal curves. 4

Throughout this chapter, we present formulas such as Equation 9.5 for sums of squares and other quantities that are used in statistics. However, we do not recommend that you use the formulas in this form to make your calculations. The reason is that these formulas are very susceptible to small round-off errors, which accumulate in a large sum (Press et al. 1986). Matrix multiplication is a much more reliable way to get the statistical solutions; in fact, it is the only way to solve more complex problems such as multiple regression. It is important for you to study these equations so that you understand how the statistics work, and it is a great idea to use spreadsheet software to try a few simple examples “by hand.” However, for analysis and publication, you should let a dedicated statistics package do the heavy numerical lifting.

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2.5 dj log10(Number of species)

244

2.0 x + 1.5

di 1.0

0.5 –1

1 2 log10(Island area)

Figure 9.3 The residual sum of squares is found by summing the squared deviation (di’s) each observation from the fitted regression line. The least-squares parameter estimates ensure that the fitted regression line–minimizes this residual – sum of squares. The + marks the midpoint of the data (X, Y ) . This regression line describes the relationship between the logarithm of island area and the logarithm of species number for plant species of the Galápagos Islands; data from Table 8.2. The regression equation is log10(Species) = 1.320 + log10(Area) × 0.331; r 2 = 0.584.

– – We could fit the regression line through (X , Y ) “by eye” and then tinker with it until we found the slope and intercept values that gave us the smallest value of RSS. Fortunately, there is an easier way to obtain the estimates of β0 and β1 that minimize RSS. But first we have to take a brief detour to discuss variances and covariances.

Variances and Covariances In Chapter 3, we introduced you to the sum of squares (SSY) of a variable, n

SSY = ∑(Yi − Y )2

(9.6)

i=1

which measures the squared deviation of each observation from the mean of the observations. Dividing this sum by (n – 1) gives the familiar formula for the sample variance of a variable: sY2 =

1 n (Yi − Y )2 n −1 ∑ i=1

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(9.7)

Variances and Covariances

By removing the exponent from Equation 9.6 and expanding the squared term, we can re-write it as n

SSY =

∑(Yi − Y )(Yi − Y )

(9.8)

i =1

Now let’s consider the situation with two variables X and Y. Instead of the sum of squares for one variable, we can define the sum of cross products (SSXY) as SS XY =

∑(X i − X)(Yi − Y )

(9.9)

i =1

and the sample covariance (sXY) as sxy =

1 n ( X i − X )(Yi − Y ) n −1 ∑ i =1

(9.10)

As we saw in Chapter 3, the sample variance is always a non-negative number. Because the square of the difference of each observation from its mean, – (Yi – Y )2, is always greater than zero, the sum of all of these squared differences must also be greater than zero. But the same is not true for the sample covariance. Suppose that relatively large values of X are consistently paired with relatively small values of Y. The first term in Equation 9.9 or Equation – 9.10 will be positive for the Xi values that are greater than X . But the second term (and hence the product) will be negative because the relatively small Yi – values are less than Y . Similarly, the relatively small Xi values (those less than – – X ) will be paired with the relatively large values of Yi (those greater than Y ). If there are lots of pairs of data organized this way, the sample covariance will be a negative number. On the other hand, if the large values of X are always associated with the large values of Y, the summation terms in Equations 9.9 and 9.10 all will be positive and will generate a large positive covariance term. Finally, suppose that X and Y are unrelated to one another, so that large or small values of X may sometimes be associated with large or small values of Y. This will generate a heterogeneous collection of covariance terms, with both positive and negative signs. The sum of these terms may be close to zero. For the Galápagos plant data, SSXY = 6.558, and sXY = 0.410. All of the elements of this covariance term are positive, except for the contributions from the islands Duncan (–0.057) and Bindlow (–0.080). Intuitively, it would seem that this measure of covariance should be related to the slope of the regression line,

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because it describes the relationship (positive or negative) between variation in the X variable and variation in the Y variable.5

Least-Squares Parameter Estimates Having defined the covariance, we can now estimate the regression parameters that minimize the residual sum of squares: s SS = XY βˆ 1 = XY 2 SSX sX

(9.11)

where the sums of squares of X is n

SS X =

∑(Xi – X )

i =1

We use the symbol βˆ 1 to designate our estimate of the slope, and to distinguish it from β1, the true value of the parameter. Keep in mind that β1 has a true value only in a frequentist statistical framework; in a Bayesian analysis, the parameters themselves are viewed as a random sample from a distribution (see Chapter 5). Later in this chapter we discuss Bayesian parameter estimation in regression models. Equation 9.11 illustrates the relationship between the slope of a regression model and the covariance between X and Y. Indeed, the slope is the covariance 5

The covariance is a single number that expresses a relationship between a single pair of variables. Suppose we have a set of n variables. For each unique pair (Xi ,Xj) of variables we could calculate each of the covariance terms sij. There are exactly n(n – 1)/2 such unique pairs, which can be determined using the binomial coefficient from Chapter 2 (see also Footnote 4 in Chapter 2). These covariance terms can then be arranged in a square n × n variance-covariance matrix, in which all of the variables are represented in each row and in each column of the matrix. The matrix entry row i, column j is simply sij. The diagonal elements of this matrix are the variances of each variable sii = si2. The matrix elements above and below the diagonal will be mirror images of one another because, for any pair of variables Xi and Xj , Equation 9.10 shows that sij = sji . The variance-covariance matrix is a key piece of machinery that is used to obtain least-squares solutions for many statistical problems (see also Chapter 12 and the Appendix). The variance-covariance matrix also makes an appearance in community ecology. Suppose that each variable represents the abundance of a species in a community. Intuitively, the covariance in abundance should reflect the kinds of interactions that occur between species pairs (negative for predation and competition, positive for mutualism, 0 for weak or neutral interactions). The community matrix (Levins 1968) and other measures of pairwise interaction coefficients can be computed from the variance-covariance matrix of abundances and used to predict the dynamics and stability of the entire assemblage (Laska and Wootton 1998).

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Least-Squares Parameter Estimates

of X and Y, scaled to the variance in X. Because the denominator (n – 1) is identical for the calculation of sXY and sX2, βˆ 1 can also be expressed as a ratio of the sum of the cross products (SSXY) to the sum of squares of X (SSX). For the Galáˆ = 0.410/1.240= 0.331. Remempagos plant data, sXY = 0.410, and sX2 = 1.240, so β 1 ber that a slope is always expressed in units of ΔY/ΔX, which in this case is the change in log10(species number) divided by the change in log10(island area). To solve for the intercept of the equation (βˆ 0), we take advantage of the fact – – that the regression line passes through the point (X, Y ). Combining this with our estimate of βˆ 1, we have βˆ 0 = Y − βˆ 1 X

(9.12)

For the Galápagos plant data, βˆ 0 = 1.867 – (0.331)(1.654) = 1.319. The units of the intercept are the same as the units of the Y variable, which in this case is log10(species number). The intercept tells you the estimate of the response variable when the value of the X variable equals zero. For our island data, X = 0 corresponds to an area of 1 square kilometer (remember that log10(1.0) = 0.0), with an estimate of 101.319 = 20.844 species. We still have one last parameter to estimate. Remember from Equation 9.3 that our regression model includes not only an intercept (βˆ 0) and a slope (βˆ 1), but also an error term (εi). This error term has a normal distribution with a mean of 0 and a variance of σ2. How can we estimate σ2? First, notice that if σ2 is relatively large, the observed data should be widely scattered about the regression line. Sometimes the random variable will be positive, pushing the datum Yi above the regression line, and sometimes it will be negative, pushing Yi below the line. The smaller σ2, the more tightly the data will cluster around the fitted regression line. Finally, if σ2 = 0, there should be no scatter at all, and the data will “toe the line” perfectly. This description sounds very similar to the explanation for the residual sum of squares (RSS), which measures the squared deviation of each observation from its fitted value (see Equation 9.5). In simple summary statistics (see Chapter 3), the sample variance of a variable measures the average deviation of each observation from the mean (see Equation 3.9). Similarly, our estimate of the variance of the regression error is the average deviation of each observation from the fitted value: n

RSS = σˆ 2 = n−2

∑ (Yi − Yˆi )2 i=1

n−2

∑ [Yi − (βˆ 0 + βˆ 1X i )] n

i=1

n−2

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(9.13)

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The expanded forms are shown to remind you of the calculation of RSS and Yˆi. As before, remember that βˆ 0 and βˆ 1 are the fitted regression parameters, and Yˆi is the predicted value from the regression equation. The square root of Equation 9.13, σˆ , is often called the standard error of regression. Notice that the denominator of the estimated variance is (n – 2), whereas we previously used (n – 1) as the denominator for the sample variance calculation (see Equation 3.9). The reason we use (n – 2) is that the denominator is the degrees of freedom, the number of independent pieces of information that we have to estimate that variance (see Chapter 3). In this case, we have already used up two degrees of freedom to estimate the intercept and the slope of the regression line. For the Galápagos plant data, σˆ = 0.320.

Variance Components and the Coefficient of Determination A fundamental technique in parametric analysis is to partition a sum of squares into different components or sources. We will introduce the idea here, and return to it again in Chapter 10. Starting with the raw data, we will consider the sum of squares of the Y variable (SSY) to represent the total variation that we are trying to partition. One component of that variation is pure or random error. This variation cannot be attributed to any particular source, other than random sampling from a normal distribution. In Equation 9.3 this source of variation is εi, and we have already seen how to estimate this residual variation by calculating the residual sums of squares (RSS). The remaining variation in Yi is not random, but systematic. Some values of Yi are large because they are associated with large values of Xi. The source of this variation is the regression relationship Yi = β0 + β1Xi. By subtraction, it follows that the remaining component of variation that can be attributed to the regression model (SSreg) is SSreg = SSY – RSS

(9.14)

By rearranging Equation 9.14, the total variation in the data can be additively partitioned into components of the regression (SSreg) and of the residuals (RSS): SSY = SSreg + RSS

(9.15)

For the Galápagos data, SSY = 3.708 and RSS = 1.540. Therefore, SSreg = 3.708 – 1.540 = 2.168. We can imagine two extremes in this “slicing of the variance pie.” Suppose that all of the data points fell perfectly on the regression line, so that any value

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of Yi could be predicted exactly knowing the value of Xi . In this case, RSS = 0, and SSY = SSreg. In other words, all of the variation in the data can be attributed to the regression and there is no component of random error. At the other extreme, suppose that the X variable had no effect on the resulting Y variable. If there is no influence of X on Y, then β1 = 0, and there is no slope to the line: Yi = β0 + εi

(9.16)

Remember that εi is a random variable with a mean of 0 and a variance of σ2. Taking advantage of the shift operation (see Chapter 2) we have Y ~ N(β0, σ)

(9.17)

In words, Equation 9.17 says, “Y is a random normal variable, with a mean (or expectation) of β0 and a standard deviation of σ.” If none of the variation in Yi can be attributed to the regression, the slope equals 0, and the Yi’s are drawn from a normal distribution with mean equal to the intercept of the regression (β0), and a variance of σ2. In this case SSY = RSS, and therefore SSreg = 0.0. Between the two extremes of SSreg = 0 and RSS = 0 lies the reality of most data sets, which reflects both random and systematic variation. A natural index that describes the relative importance of regression versus residual variation is the familiar r 2, or coefficient of determination: r2 =

SSreg SSY

SSreg SSreg + RSS

(9.18)

The coefficient of determination tells you the proportion of the variation in the Y variable that can be attributed to variation in the X variable through a simple linear regression. This proportion varies from 0.0 to 1.0. The larger the value, the smaller the error variance and the more closely the data match the fitted regression line. For the Galápagos data, r 2 = 0.585, about midway between no correlation and a perfect fit. If you convert the r 2 value to a scale of 0 to 100, it is often described as the percentage of variation in Y “explained” by the regression on X. Remember, however, that the causal relationship between the X and Y variable is an hypothesis that is explicitly proposed by the investigator (see Equation 9.1). The coefficient of determination—no matter how large—does not by itself confirm a cause-and-effect relationship between two variables. A related statistic is the product-moment correlation coefficient, r. As you might guess, r is just the square root of r 2. However, the sign of r (positive or

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negative) is determined by the sign of the regression slope; it is negative if β1 < 0 and positive if β1 > 0. Equivalently, r can be calculated as r=

SS XY (SS X )(SSY )

sXY sX sY

(9.19)

with the positive or negative sign coming from the sum of cross products term in the numerator.6

Hypothesis Tests with Regression So far, we have learned how to fit a straight line to continuous X and Y data, and how to use the least-squares criterion to estimate the slope, the intercept, and the variance of the fitted regression line. The next step is to test hypotheses about the fitted regression line. Remember that the least squares calculations give us only estimates (βˆ 0, βˆ 1, σˆ 2) of the true values of the parameters (β0, β1, σ2). Because there is uncertainty in these estimates, we want to test whether some of these parameter estimates differ significantly from zero. In particular, the underlying assumption of cause and effect is embodied in the slope parameter. Remember that, in setting up the regression model, we assume that X causes Y (see Figure 6.2 for other possibilities). The magnitude of β1 measures the strength of the response of Y to changes in X. Our null hypothesis is that β1 does not differ from zero. If we cannot reject this null hypothesis, there is no compelling evidence of a functional relationship between the X and the Y variables. Framing the null and alternative hypotheses in terms of our models, we have Yi = β0 + εi (Null hypothesis)

(9.20)

Yi = β0 + β1Xi + εi (Alternative hypothesis)

(9.21)

Rearranging Equation 9.19 reveals a close connection between r and β1, the slope of the regression: ⎛s ⎞ β1 = r ⎜ Y ⎟ ⎝ sX ⎠

Thus, the slope of the regression line turns out to be the correlation coefficient “rescaled” to the relative standard deviations of Y and X.

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The Anatomy of an ANOVA Table

This null hypothesis can be tested by first organizing the data into an analysis of variance (ANOVA) table. Although an ANOVA table is naturally associated with an analysis of variance (see Chapter 10), the partitioning of the sum of squares is common to ANOVA, regression, and many other generalized linear models (McCullagh and Nelder 1989). Table 9.1 illustrates a complete ANOVA table with all of its components and equations. Table 9.2 illustrates the same ANOVA table with calculations for the Galápagos plant data. The abbreviated form of Table 9.2 is typical for a scientific publication. The ANOVA table has a number of columns that summarize the partitioning of the sum of squares. The first column is usually labeled Source, meaning the component or source of variation. In the regression model, there are only two sources: the regression and the error. We have also added a third source, the total, to remind you that the total sum of squares equals the sum of the regression and the error sums of squares. However, the row giving the total sums of squares usually is omitted from published ANOVA tables. Our simple regression model has only two sources of variation, but more complex models may have several sources of variation listed.

TABLE 9.1 Complete ANOVA table for single factor linear regression

Source

Mean Degrees of square freedom (df) Sum of squares (SS) (MS) n

Regression

SSreg = ∑ (Yˆi − Y )2 i=1

Residual

n–2

Total

n–1

RSS = ∑ (Yi − Yˆi )2 i=1 n

SSY = ∑(Yi − Y )2 i =1

SSreg 1 RSS (n − 2) SSY (n −1)

Expected mean square n

σ 2 + β12 ∑ X i2 i =1

F-ratio

P-value

SSreg / 1

Tail of the F RSS / (n − 2) distribution with 1, n – 2 degrees of freedom

σ2 σY2

The first column gives the source of variation in the data. The second column gives the degrees of freedom (df) associated with each component. For a simple linear regression, there is only 1 degree of freedom associated with the regression, and (n – 2) associated with the residual. The degrees of freedom total to (n – 1) because 1 degree of freedom is always used to estimate the grand mean of the data. The single factor regression model partitions the total variation into a component explained by the regression and a remaining (“unexplained”) residual. The sums – of squares are calculated using the observed Y values (Yi), the mean of the Y values (Y ), and the Y values predicted ˆ by the linear regression model (Yi ). The expected mean squares are used to construct an F-ratio to test the null hypothesis that the variation associated with the slope term (β1) equals 0.0. The P-value for this test is taken from a standard table of F-values with 1 degree of freedom for the numerator and (n – 2) degrees of freedom for the denominator. These basic elements (source, df, sum of squares, mean squares, expected mean square, F-ratio, and P-value) are common to all ANOVA tables in regression and analysis of variance (see Chapter 10).

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TABLE 9.2 Publication form of ANOVA table for regression analysis of Galápagos plants species-area data Source

F-ratio

Regression Residual

1 15

2.168 1.540

2.168 0.103

21.048

0.000329

The raw data are from Table 8.2; the formulas for these calculations are given in Table 9.1. The total sum of squares and degrees of freedom are rarely included in a published ANOVA table. In many publications, these results would be reported more compactly as “The linear regression of log-species richness against log-area was highly significant (model F1,15 = 21.048, p < 0.001). Thus, the best-fitting equation was log10(Species richness) = 1.867 + 0.331 × log(island area); r 2 = 0.585.” Published regression statistics should include the coefficient of determination (r 2) and the least-squares estimators of the slope (0.331) and the intercept (1.867).

The second column gives the degrees of freedom, usually abbreviated df. As we have discussed, the degrees of freedom depend on how many pieces of independent information are available to estimate the particular sum of squares. If the sample size is n, there is 1 degree of freedom associated with the regression model (specifically the slope), and (n – 2) degrees of freedom associated with the error. The total degrees of freedom is (1 + n – 2) = (n – 1). The total is only (n – 1) – because 1 degree of freedom was used in estimating the grand mean, Y . The third column gives the sum of squares (SS) associated with particular source of variation. In the expanded Table 9.1, we showed the formulas used, but in published tables (e.g., Table 9.2), only the numerical values for the sums of squares would be given. The fourth column gives the mean square (MS), which is simply the sum of squares divided by its corresponding degrees of freedom. This division is analogous to calculating a simple variance by dividing SSY by (n – 1). The fifth column is the expected mean square. This column is not presented in published ANOVA tables, but it is very valuable because it shows exactly what is being estimated by each of the different mean squares. It is these expectations that are used to formulate hypothesis tests in ANOVA. The sixth column is the calculated F-ratio. The F-ratio is a ratio of two different mean square values. The final column gives the tail probability value corresponding to the particular F-ratio. Specifically, this is the probability of obtaining the observed F-ratio (or a larger one) if the null hypothesis is true. For the simple linear regression model, the null hypothesis is that β1 = 0, implying no functional relationship between the X and Y variables. The probability value depends on the size of the F-ratio and on the number of degrees of freedom associated with the numerator and denominator mean squares. The probability values can be looked up in sta-

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Hypothesis Tests with Regression

tistical tables, but they are usually printed as part of the standard regression output from statistics packages. In order to understand how the F-ratio is constructed, we need to examine the expected mean square values. The expected value of the regression mean square is the sum of the regression error variance and a term that measures the regression slope effect: n

E( MSreg) = σ 2 + β12 ∑ X i2

(9.22)

i =1

In contrast, the expected value of the residual mean square is simply the regression error variance: E(MSresid) = σ2

(9.23)

Now we can understand the logic behind the construction of the F-ratio. The F-ratio for the regression test uses the regression mean square in the numerator and the residual mean square in the denominator. If the true regression slope is zero, the second term in Equation 9.22 (the numerator of the F-ratio) also will equal zero. As a consequence, Equation 9.22 (the numerator of the Fratio) and Equation 9.23 (the denominator of the F-ratio) will be equal. In other words, if the slope of the regression (β1) equals zero, the expected value of the F-ratio will be 1.0. For a given amount of error variance, the steeper the regression slope, the larger the F-ratio. Also, for a given slope, the smaller the error variance, the larger the F-ratio. This also makes intuitive sense because the smaller the error variance, the more tightly clustered the data are around the fitted regression line. The interpretation of the P-value follows along the lines developed in Chapter 4. The larger the F-ratio (for a given sample size and model), the smaller the P-value. The smaller the P-value, the more unlikely it is that the observed Fratio would have been found if the null hypothesis were true. With P-values less than the standard 0.05 cutoff, we reject the null hypothesis and conclude that the regression model explains more variation than could be accounted for by chance alone. For the Galápagos data, the F-ratio = 21.048. The numerator of this F-ratio is 21 times larger than the denominator, so the variance accounted for by the regression is much larger than the residual variance. The corresponding P-value = 0.0003. Other Tests and Confidence Intervals

As you might suspect, all hypothesis tests and confidence intervals for regression models depend on the variance of the regression (σˆ 2). From this, we can

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calculate other variances and significance tests (Weisberg 1980). For example, the variance of the estimated intercept is ⎛ 1 X2 ⎞ σˆ β2ˆ = σˆ 2 ⎜ + 0 ⎝ n SSX ⎟⎠

(9.24)

An F-ratio also can be constructed from this variance to test the null hypothesis that β0 = 0.0. Notice that the intercept of the regression line is subtly different from the intercept that is calculated when the model has a slope of zero (Equation 9.16): Yi = β0 + εi If the model has a slope of 0, the expected value of the intercept is simply the average of the Yi values: – E(β0) = Y However, for the regression model with a slope term, the intercept is the expected value when Xi = 0, that is, E (β 0 ) = Yˆi | ( X i = 0) A simple 95% confidence interval can be calculated for the intercept as

βˆ0 − t(α ,n− 2)σˆ βˆ ≤ β0 ≤ βˆ0 + t(α , n− 2)σˆ βˆ 0

(9.25)

where α is the (two-tailed) probability level (α = 0.025 for a 95% confidence interval), n is the sample size, t is the table value from the t-distribution for the specified α and n, and σˆ βˆ is calculated as the square root of Equation 9.24. 0 Similarly, the variance of the slope estimator is

σˆ 2 σˆ β2ˆ = 1 SSX

(9.26)

and the corresponding confidence interval for the slope is

βˆ1 − t(α ,n− 2)σˆ βˆ ≤ β1 ≤ βˆ1 + t(α ,n− 2)σˆ βˆ 1

(9.27)

For the Galápagos data, the 95% confidence interval for the intercept (β0) is 1.017 to 1.623, and the 95% confidence interval for the slope (β1) is 0.177 to 0.484 (remember these are log10 values). Because neither of these confidence intervals bracket 0.0, the corresponding F-tests would cause us to reject the null

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hypothesis that both β0 and β1 equal 0. If β1 equals 0, the regression line is horizontal, and the dependent variable [log10(species number)] does not systematically increase with changes in the independent variable [log10(island area)]. If β0 equals 0, the dependent variable takes on a value of 0 when the independent variable is 0. Although the null hypothesis has been rejected in this case, the observed data do not fall perfectly in a straight line, so there is going to be uncertainty associated with any particular value of the X variable, For example, if you were to repeatedly sample different islands that were identical in island area (X), there would be variation among them in the (log-transformed) number of species recorded.7 The variance of the fitted value Yˆ is ⎛ 1 ( X − X )2 ⎞ sˆ 2(Yˆ |X ) = sˆ 2 ⎜ + i SS X ⎟⎠ ⎝n

(9.28)

and a 95% confidence interval is Yˆ − t ( a ,n−2)sˆ (Yˆ |X ) ≤ Yˆ ≤ Yˆ + t (a ,n−2)sˆ (Yˆ |X )

(9.29)

This confidence interval does not form a parallel band that brackets the regression line (Figure 9.4). Rather, the confidence interval flares out the farther – – away we move from X because of the term (Xi – X )2 in the numerator of Equation 9.28. This widening confidence interval makes intuitive sense. The closer we are to the center of the cloud of points, the more confidence we have in – estimating Y from repeated samples of X. In fact, if we choose X as the fitted 7

Unfortunately, there is a mismatch between the statistical framework of random sampling and the nature of our data. After all, there is only one Galápagos archipelago, which has evolved a unique flora and fauna, and there are no multiple replicate islands that are identical in area. It seems dubious to treat these islands as samples from some larger sample space, which itself is not clearly defined—would it be volcanic islands? tropical Pacific islands? isolated oceanic islands? We could think of the data as a sample from the Galápagos, except that the sample is hardly random, as it consists of all the large major islands in the archipelago. There are a few additional islands from which we could collect data from, but these are considerably smaller, with very few species of plants and animals. Some islands are so small as to be essentially empty, and these 0 data (for which we cannot simply take logarithms) have important effects on the shape of the species–area relationship (Williams 1996). The fitted regression line for all islands may not necessarily be the same as for sets of large versus small islands. These problems are not unique to species–area data. In any sampling study, we must struggle with the fact that the sample space is not clearly defined and the replicates we collect may be neither random nor independent of one another, even though we use statistics that rely on these assumptions.

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Figure 9.4 Regression line (dark blue), 95% confidence interval (light blue lines), and 95% prediction interval (gray lines) for a log-log regression of Galápagos plant species number against island area (see Table 8.2). The confidence interval describes the uncertainty in the collected data, whereas the prediction interval is used to evaluate new data not yet collected or included in the analysis. Notice that the confidence intervals flare outward as we move further away from the average island area in the collected data. The flaring confidence intervals reflect increasing uncertainty about predictions for X variables that are very different from those already collected. Notice also that these intervals are described on a logarithmic scale. Back-transformation (eY ) is necessary to convert the units to numbers of species. Back-transformed confidence intervals are very wide and asymmetric around the predicted value.

3.5 log10(Number of species)

256

3.0 2.5 2.0 1.5 1.0 0.5 0 –1

log10(Island area)

value, the standard deviation of the fitted value is equivalent to a standard error of the average fitted value (see Chapter 3): σˆ (Yˆ |X ) = σˆ / n The variance is minimized here because there are observed data both above and – below the fitted value. However, as we move away from X, there are fewer data in the neighborhood, so the prediction becomes more unreliable. A useful distinction can be made here between interpolation and extrapolation. Interpolation is the estimation of new values that are within the range of the data we have collected, whereas extrapolation means estimating new values beyond the range of data (see Figure 9.2). Equation 9.29 ensures that confidence intervals for fitted values always will be smaller for interpolation than for extrapolation. Finally, suppose we discover a new island in the archipelago, or sample an island that was not included in our original census. This new observation is designated ( X˜ , Y˜)

We will use the fitted regression line to construct a prediction interval for the new value, which is subtly different from constructing a confidence interval for a fitted value. The confidence interval brackets uncertainty about an estimate based on the available data, whereas the prediction interval brackets uncertainty about an estimate based on new data. The variance of the prediction for a single fitted value is ⎛ 1 ( X˜ − X )2 ⎞ s 2(Y˜|X˜ ) = sˆ 2 ⎜1 + + SS X ⎟⎠ ⎝ n

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(9.30)

Assumptions of Regression

and the corresponding prediction interval is Y˜ − t (α,n−2)σˆ (Y˜|X˜ ) ≤ Y˜ ≤ Y˜ + t (α,n−2)σˆ (Y˜|X˜ )

(9.31)

This variance (Equation 9.30) is larger than the variance for the fitted value (Equation 9.28). Equation 9.30 includes variability both from the error associated with the new observation, and from the uncertainty in the estimates of the regression parameters from the earlier data.8 To close this section, we note that it is very easy (and seductive) to make a point prediction from a regression line. However, for most kinds of ecological data, the uncertainty associated with that prediction usually is too large to be useful. For example, if we were to propose a nature reserve of 10 square kilometers based only on the information contained in the Galápagos species–area relationship, the point prediction from the regression line is 45 species. However, the 95% prediction interval for this forecast is 9 to 229 species. If we increased the area 10-fold to 100 square kilometers, the point prediction is 96 species, with a range of 19 to 485 species (both ranges based on a back-transformed prediction interval from Equation 9.31).9 These wide prediction intervals not only are typical for species–area data but also are typical for most ecological data.

Assumptions of Regression The linear regression model that we have developed relies on four assumptions: 1. The linear model correctly describes the functional relationship between X and Y. This is the fundamental assumption. Even if the overall relationship is non-linear, a linear model may still be appropriate over a limited range of the X variable (see Figure 9.2). If the assumption of linearity is violated, the estimate of σ2 will be inflated because it will include both a random error and a fixed error component; the latter represents the difference between the true function and the linear one that has been fit to the data. And, if the true relationship is not linear, 8

A final refinement is that Equation 9.31 is appropriate only for a single prediction. If multiple predictions are made, the α value must be adjusted to create a slightly broader simultaneous prediction interval. Formulas also exist for inverse prediction intervals, in which a range of X values is obtained for a single value of the Y variable (see Weisberg 1980).

9 To be picky, a back-transformed point estimates the median, not the mean, of the distribution. However, back-transformed confidence intervals are not biased because the quantiles (percentage points) translate across transformed scales.

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predictions derived from the model will be misleading, particularly when extrapolated beyond the range of the data. 2. The X variable is measured without error. This assumption allows us to isolate the error component entirely as random variation associated with the response variable (Y). If there is error in the X variable, the estimates of the slope and intercept will be biased. By assuming there is no error in the X variable, we can use the least-squares estimators, which minimize the vertical distance between each observation and its predicted value (the di’s in Figure 9.3). With errors in both the X and Y variable, one strategy would be to minimize the perpendicular distance between each observation and the regression line. This so-called Model II regression is commonly used in principle components and other multivariate techniques (see Chapter 12). However, because the leastsquares solutions have proven so efficient and are used so commonly, this assumption often is quietly ignored.10 3. For any given value of X, the sampled Y values are independent with normally distributed errors. The assumption of normality allows us to use parametric theory to construct confidence intervals and hypothesis tests based on the F-ratio. Independence, of course, is the critical assumption for all sample data (see Chapter 6), even though it often is violated to an unknown degree in observational studies. If you suspect that the value Yi influences the next observation you collect (Yi+1), a time-series analysis may remove the correlated components of the error variation. Time-series analysis is discussed briefly in Chapter 6. 4. Variances are constant along the regression line. This assumption allows us to use a single constant σ2 for the variance of regression line. If variances depend on X, then we would require a variance function, or an entire family of variances, each predicated on a particular value of X. Non-constant variances are a common problem in regression that can be recognized through diagnostic plots (see the following section) and sometimes remedied through transformations of the original X or Y variables (see Chapter 8). However, there is no guarantee that a transformation will linearize the relationship and generate constant variances. In such cases, other methods (such as non-linear regression, described

10 Although it is rarely mentioned, we make a similar assumption of no errors for the categorical X variable in an analysis of variance (see Chapter 10). In ANOVA, we have to assume that individuals are correctly assigned to groups (e.g., no errors in species identification) and that all individuals within a group have received an identical treatment (e.g., all replicates in a “low pH” treatment received exactly the same pH level).

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later in this chapter) or generalized linear models (McCullagh and Nelder 1989) should be used. If all four of these assumptions are met, the least-squares method provides unbiased estimators of all the model parameters. They are unbiased because repeated samples from the same population will yield slope and intercept estimates that, on average, are the same as the true underlying slope and intercept values for the population.

Diagnostic Tests For Regression We have now seen how to obtain least-squares estimates of linear regression parameters and how to test hypotheses about those parameters and construct appropriate confidence intervals. However, a regression line can be forced through any set of {X,Y} data, regardless of whether the linear model is appropriate. In this section, we present some diagnostic tools for determining how well the estimated regression line fits the data. Indirectly, these diagnostics also help you to evaluate the extent to which the data meet the assumptions of the model. The most important tool in diagnostic analysis is the set of residuals, {di}, which represents the differences between the observed values (Yi) and the values predicted by the regression model (Yˆi ) of Equation 9.4. The residuals are used to estimate the regression variance, but they also provide important information about the fit of the model to the data. Plotting Residuals

Perhaps the single most important graph for diagnostic analysis of the regression model is the plot of the residuals (di) versus the fitted values (Yˆi ). If the linear model fits the data well, this residual plot should exhibit a scatter of points that approximately follow a normal distribution and are completely uncorrelated with the fitted values (Figure 9.5A). In Chapter 11, we will explain the Kolmogorov-Smirnov test, which can be used to formally compare the residuals to a normal distribution (see Figure 11.4). Two kinds of problems can be spotted in residual plots. First, if the residuals themselves are correlated with the fitted values, it means that the relationship is not really linear. The model may be systematically over-estimating or underestimating Yˆi for high values of the X variable (Figure 9.5B). This can happen, for example, when a straight line is forced through data that really represent an asymptotic, logarithmic, or other non-linear relationship. If the residuals first rise above the fitted values then fall below, and then rise above again, the data

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Figure 9.5 Hypothetical patterns for diagnostic plots of residuals (di) versus fitted values (Yˆi) in linear regression. (A) Expected distribution of residuals for a linear model with a normal distribution of errors. If the data are well fit by the linear model, this is the pattern that should be found in the residuals. (B) Residuals for a non-linear fit; here the model systemically overestimates the actual Y value as X increases. A mathematical transformation (e.g., logarithm, square root, or reciprocal) may yield a more linear relationship. (C) Residuals for a quadratic or polynomial relationship. In this case, large positive residuals occur for very small and very large values of the X variable. A polynomial transformation of the X variable (X2 or some higher power of X) may yield a linear fit. (D) Residuals with heteroscedascity (increasing variance). In this case, the residuals are neither consistently positive nor negative, indicating that the fit of the model is linear. However, the average size of the residual increases with X (heteroscedascity), suggesting that measurement errors may be proportional to the size of the X variable. A logarithmic or square root transformation may correct this problem. Transformations are not a panacea for regression analyses and do not always result in linear relationships.

may indicate a quadratic rather than a linear relationship (Figure 9.5C). Finally, if the residuals display an increasing or decreasing funnel of points when plotted against the fitted values (Figure 9.5D), the variance is heteroscedastic: either increasing or decreasing with the fitted values. Appropriate data transformations, discussed in Chapter 8, can address some or all of these problems. Resid-

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(A)

Figure 9.6 Residual plot for species–area relationship for plants of Galápagos Islands (see Table 8.2). In a residual plot, the x-axis is the predicted value from the regression equation (Yˆi), and the y-axis is the residual, which is the difference between the observed and fitted values. In a dataset that matches the predictions of the regression model, the residual plot should be a normally distributed cloud of points centered on the average value of 0 (see Figure 9.5a). (A) Residual plot for regression of untransformed data (see Figure 8.5). There are too many negative residuals at small fitted values, and the distribution of residuals does not appear normal. (B) Residual plot for regression of the same data after a log10 transformation of both the x- and y-axes (see Figures 8.6 and 9.3). The transformation has considerably improved the distribution of the residuals, which no longer deviate systematically at large or small fitted values. Chapter 11 describes the KolmogorovSmirnov test, which can be used to test the residuals for deviations from a normal distribution (see Figure 11.4).

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ual plots also may highlight outliers, points that fall much further from the regression prediction than most of the other data.11 To see the effects of transformations, compare Figures 9.6A and 9.6B, which illustrate residual plots for the Galápagos data before and after log10 transformation. Without the transformation, there are too many negative residuals, and they are all clustered around very small values of Yˆi (Figure 9.6A). After trans-

Standardized residuals (also called studentized residuals) also are calculated by many statistics packages. Standardized residuals are scaled to account for the distance of each point from the center of the data. If the residuals are not standardized, data – points far away from X appear to be relatively well-fit by the regression, whereas points closer to the center of the data cloud appear to not fit as well. Standardized residuals also can be used to test whether particular observations are statistically significant outliers. Other residual distance measures include Cook’s distance and leverage values. See Sokal and Rohlf (1995) for details.

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formation, the positive and negative residuals are evenly distributed and not associated with Yˆi (Figure 9.6B). Other Diagnostic Plots

Residuals can be plotted not only against fitted values, but also against other variables that might have been measured. The idea is to see whether there is any variation lurking in the errors that can be attributed to a systematic source. For example, we could plot the residuals from Figure 9.6B against some measure of habitat diversity on each island. If the residuals were positively correlated with habitat diversity, then islands that have more species than expected on the basis of area usually have higher habitat diversity. In fact, this additional source of systematic variation can be included in a more complex multiple regression model, in which we fit coefficients for two or more predictor variables. Plotting residuals against other predictor variables is often a more simple and reliable exploratory tactic than assuming a model with linear effects and linear interactions. It also may be informative to plot the residuals against time or data collection order. This plot may indicate altered measurement conditions during the time period that the data were collected, such as increasing temperatures through the course of the day in an insect behavior study, or a pH meter with a weakening battery that gives biased results for later measurements. These plots again remind us of the surprising problems that can arise by not using randomization when we are collecting data. If we collect all of our behavior measurements on one insect species in the morning, or measure pH first in all of the control plots, we have introduced an unexpected confounding source of variation in our data. The Influence Function

Residual plots do a good job of revealing non-linearity, heteroscedascity, and outliers. However, potentially more insidious are influential data points. These data may not show up as outliers, but they do have an inordinate influence on the slope and intercept estimates. In the worst situation, influential data points that are far removed from the cloud of typical data points can completely “swing” a regression line and dominate the slope estimate (see Figure 7.3). The best way to detect such points is to plot an influence function. The idea is simple, and is actually a form of statistical jackknifing (see Footnote 2 in Chapter 5; see also Chapter 12). Take the first of the n replicates in your dataset and delete it from the analysis. Recompute the slope, intercept, and probability value. Now replace that first data point and remove the second data point. Again calculate the regression statistics, replace the datum, and continue through the data list. If you have n original data points, you will end up with n different regression analyses, each of which is based on a total of (n – 1) points. Now take the slope

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and intercept estimates for each of those analyses and plot them together in a single scatter plot (Figure 9.7). On this same plot, graph the slope and intercept estimates from the complete data set. This graph of intercept versus slope estimates will itself always have a negative slope because as the regression line tips up, the slope increases and the intercept decreases. It may also be informative to create a histogram of r 2 values or tail probabilities that are associated with each of those regression models. The influence function illustrates how much your estimated regression parameters would have changed just by the exclusion of a single datum. Ideally, the jackknifed parameter estimates should cluster tightly around the estimates from the full dataset. A cluster of points would suggest that the slope and intercept values are stable and would not change greatly with the deletion (or addition) of a single data point. On the other hand, if one of the points is very distant from the cluster, the slope and intercept are highly influenced by that sin(A) 105 100 95 βˆ 0

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Figure 9.7 Influence function for species-area linear regression of Galápagos island plant data (see Table 8.2). In an influence function plot, each point represents the recomputed slope (βˆ 1) and intercept (βˆ 0) following deletion of a single observation from the dataset. The blue point is the slope and intercept estimate for the complete data set. (A) In the regression model with untransformed data, the observation for the large island Albemarle (see Figure 8.5) has a very large influence on the slope and intercept, generating the extreme point in the lower right-hand corner of the graph. (B) Influence function calculated for regression line fitted to log10 transformed data. After the logarithmic transformation (see Figures 8.6 and 9.3), there is now a more hom*ogenous cloud of points in the influence function. Although the slope and intercept will change following the deletion of each observation, no single data point has an extreme influence on the calculated slope and intercept. In this dataset, the logarithmic transformation not only improves the linear fit of the data, but it also stabilizes the slope and intercept estimates, so they are not dominated by one or two influential data points.

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gle datum, and we should be careful about what conclusions we draw from this analysis. Similarly, when we examine the histogram of probability values, we want to see all of the observations clustered nicely around the P-value that was estimated for the full dataset. Although we expect to see some variation, we will be especially troubled if one of the jackknifed samples yields a P-value very different from all the rest. In such a case, rejecting or failing to reject the null hypothesis hinges on a single observation, which is especially dangerous. For the Galápagos data, the influence function highlights the importance of using an appropriate data transformation. For the untransformed data, the largest island in the dataset dominates the estimate of β1. If this point is deleted, the slope increases from 0.035 to 0.114 (Figure 9.7A). In contrast, the influence function for the transformed data shows much more consistency, although βˆ 1 still ranged from 0.286 to 0.390 following deletion of a single datum (Figure 9.7B). For both data sets, probability values were stable and never increased above P = 0.023 for any of the jackknife analyses. Using the jackknife in this way is not unique to regression. Any statistical analysis can be repeated with each datum systematically deleted. It can be a bit time-consuming,12 but it is an excellent way to assess the stability and general validity of your conclusions.

Monte Carlo and Bayesian Analyses The least-squares estimates and hypotheses tests are representative of classic frequentist parametric analysis, because they assume (in part) that the error term in the regression model is a normal random variable; that the parameters have true, fixed values to be estimated; and that the P-values are derived from probability distributions based on infinitely large samples. As we emphasized in Chapter 5, Monte Carlo and Bayesian approaches differ in philosophy or implementation (or both), but they also can be used for regression analysis. Linear Regression Using Monte Carlo Methods

For the Monte Carlo approach, we still use all of the least-squares estimators of the model parameters. However, in a Monte Carlo analysis, we relax the assumption of normally distributed error terms, so that significance tests are no longer based on comparisons with F-ratios. Instead, we randomize and resample the data and simulate the parameters directly to estimate their distribution. 12

Actually, there are clever matrix solutions to obtaining the jackknifed values, although these are not implemented in most computer packages. The influence function is most important for small data sets that can be analyzed quickly by using case selection options that are available in most statistical software.

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Monte Carlo and Bayesian Analyses

For linear regression analysis, what is an appropriate randomization method? We can simply reshuffle the observed Yi values and pair them randomly with one of the Xi values. For such a randomized dataset, we can then calculate the slope, intercept, r 2, or any other regression statistics using the methods described in the previous section. The randomization method effectively breaks up any covariation between the X and Y variables and represents the null hypothesis that the X variable (here, island area) has no effect on the Y variable (here, species richness). Intuitively, the expected slope and r 2 values should be approximately zero for the randomized data sets, although some values will differ by chance. For the Galápagos data, 5000 reshufflings of the data generated a set of slope values with a mean of –0.002 (very close to 0), and a range of –0.321 to 0.337 (Figure 9.8). However, only one of the simulated values (0.337) exceeded the observed slope of 0.331. Therefore, the estimated tail probability is 1/5000 =

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Figure 9.8 Monte Carlo analysis of slope values for the log-log transformed species–area relationship of plant species on the Galápagos Islands (see Table 8.2; see also Figures 8.6 and 9.3). Each observation in the histogram represents the leastsquares slope of a simulated dataset in which the observed X and Y values were randomly reshuffled. The histogram illustrates the distribution of 5000 such random datasets. The observed slope from the original data is plotted as an arrow on the histogram. The observed slope of 0.331 exceeded all but 1 of the 5000 simulated values. Therefore, the estimated tail probability of obtaining a result this extreme under the null hypothesis is P = 1/5000 = 0.0002. This probability estimate is consistent with the probability calculated from the F-ratio for the standard regression test of these data (P = 0.0003; see Table 9.2).

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0.0002. By comparison, the parametric analysis gave essentially the same result, with a P-value equal to 0.0003 for F1,15 = 21.118 (see Table 9.2). One interesting result from the Monte Carlo analysis is that the P-values for the intercept, slope, and r 2 tests are all identical. Naturally, the observed parameter estimates and simulated distributions for each of these variables are all very different. But why should the tail probabilities be the same in each case? The reason is that β0, β1, and r 2 are not algebraically independent of one another; the randomized values reflect that interdependence. Consequently, the tail probabilities come out the same for these fitted values when they are compared to distributions based on the same randomized datasets. This doesn’t happen in parametric analysis because P-values are not determined by reshuffling the observed data. Linear Regression Using Bayesian Methods

Our Bayesian analysis begins with the assumption that the parameters of the regression equation do not have true values to be estimated, but rather are random variables (see Chapter 5). Therefore, the estimates of the regression parameters—the intercept, slope, and error term—are reported as probability distributions, not as point values. To carry out any Bayesian analysis, including a Bayesian regression, we require an initial expectation of the shapes of these distributions. These initial expectations, called prior probability distributions, come from previous research or understanding of the study system. In the absence of previous research, we can use an uninformative prior probability distribution, as described in Chapter 5. However, in the case of the Galápagos data, we can take advantage of the history of studies on the relationship between island area and species richness that pre-dated Preston’s (1962) analysis of the Galápagos data. For this example, we will carry out the Bayesian regression analysis using data that Preston (1962) compiled for other species-area relationships in the ecological literature. He summarized the previous information available (in his Table 8), and found that, for 6 other studies, the average slope of the species-area relationship (on a log10-log10 plot) = 0.278 with a standard deviation of 0.036. Preston (1962) did not report his estimates of the intercept, but we calculated it from the data he reported: the mean intercept = 0.854 and the standard deviation = 0.091. Lastly, we need an estimate of the variance of the regression error ε, which we calculated from his data as 0.234. With this prior information, we use Bayes Theorem to calculate new distributions (the posterior probability distributions) on each of these parameters. Posterior probability distributions take into account the previous information plus the data from the Galápagos Islands. The results of this analysis are βˆ 0 ~ N(1.863, 0.157) βˆ ~ N(0.328, 0.145) 1

σˆ 2 = 0.420

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Monte Carlo and Bayesian Analyses

In words, the intercept is a normal random variable with mean = 1.863 and standard deviation 0.157, the slope is a normal random variable with mean = 0.328 and standard deviation = 0.145, and the regression variance is a normal random variable with mean 0 and variance 0.420.13 How do we interpret these results? The first thing to re-emphasize is that in a Bayesian analysis, the parameters are considered to be random variables—they do not have fixed values. Thus, the linear model (see Equation 9.3) needs to be re-expressed as Yi ~ N(β0 + β1 Xi , σ2)

(9.32)

In words, “each observation Yi is drawn from a normal distribution with mean = β0 + β1Xi and variance = σ2.” Similarly, both the slope β0 and the intercept β1 are themselves normal random variables with estimated means βˆ 0 and βˆ 1 and corresponding estimated variances sˆ02 and sˆ12. How do these estimates apply to our data? First, the regression line has a slope of 0.328 (the expected value of the β1) and an intercept of 1.863. The slope differs only slightly from the least-squares regression line shown in Figure 9.4, which has a slope of 0.331. The intercept is substantially higher than the least-squares fit, suggesting that the larger number of small islands contributed more to this regression than the few large islands. Second, we could produce a 95% credibility interval. The computation essentially is the same as that shown in Equations 9.25 and 9.27, but the interpretation of the Bayesian credibility interval is that the mean value would lie within the credibility interval 95% of the time. This differs from the interpretation of the 95% confidence interval, which is that the confidence interval will include the true value 95% of the time (see Chapter 3). Third, if we wanted to predict the species richness for a given island size Xi , the predicted number of species would be found by first drawing a value for the slope β1 from a normal distribution with mean = 0.328 and standard deviation = 0.145, and then multiplying this value by Xi. Next, we draw a value for the intercept β0 from a normal distribution with mean = 1.863 and standard

13 Alternatively, we could conduct this analysis without any prior information, as discussed in Chapter 4. This more “objective” Bayesian analysis uses uninformative prior probability distributions on the regression parameters. The results for the analysis using uninformative priors are βˆ 0 ~ N(1.866, 0.084), βˆ 1 ~ N(0.329, 0.077), and σˆ 2 = 0.119. The lack of substantive difference between these results and those based on the informed prior distribution illustrates that good data overwhelm subjective opinion.

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deviation = 0.157. Last, we add “noise” to this prediction by adding an error term drawn from a normal distribution with X = 0 and σ2 = 0.42. Finally, some words about hypothesis testing. There is no way to calculate a P-value from a Bayesian analysis. There is no P-value because we are not testing a hypothesis about how unlikely our data are given the hypothesis [as in the frequentist’s estimate of P(data|H0)] or how frequently we would expect to get these results given chance alone (as in the Monte Carlo analysis). Rather, we are estimating values for the parameters. Parametric and Monte Carlo analyses estimate parameter values too, but the primary goal often is to determine if the estimate is significantly different from 0. Frequentist analyses also can incorporate prior information, but it is still usually in the context of a binary (yes/no) hypothesis test. For example, if we have prior information that the slope of the species–area relationship, β1 = 0.26, and our Galápagos data suggest that β1 = 0.33, we can test whether the Galápagos data are “significantly different” from previously published data. If the answer is “yes,” what do you conclude? And if the answer is “no”? A Bayesian analysis assumes that you’re interested in using all the data you have to estimate the slope and intercept of the species area relationship.

Other Kinds of Regression Analyses The basic regression model that we have developed just scratches the surface of the kinds of analyses that can be done with continuous predictor and response variables. We provide only a very brief overview to other kinds of regression analyses. Entire books have been written on each of these individual topics, so we cannot hope to give more than an introduction to them. Nevertheless, many of the same assumptions, restrictions, and problems that we have described for simple linear regression apply to these methods as well. Robust Regression

The linear regression model estimates the slope, intercept, and variance by minimizing the residual sum of squares (RSS) (see Equation 9.5). If the errors follow a normal distribution, this residual sum of squares will provide unbiased estimates of the model parameters. The least-squares estimates are sensitive to outliers because they give heavier weights to large residuals. For example, a residual of 2 contributes 22 = 4 to the RSS, but a residual of 3 contributes 32 = 9 to the RSS. The penalty added to large residuals is appropriate, because these large values will be relatively rare if the errors are sampled from a normal distribution. However, when true outliers are present—aberrant data points (including erroneous data) that were not sampled from the same distribution—they can

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seriously inflate the variance estimates. Chapter 8 discussed several methods for identifying and dealing with outliers in your data. But another strategy is to fit the model using a residual function other than least-squares, one that is not so sensitive to the presence of outliers. Robust regression techniques use different mathematical functions to quantify residual variation. For example, rather than squaring each residual, we could use its absolute value: n

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As with RSS, this residual is large if the individual derivations (di) are large. However, very large deviations are not penalized as heavily because their values are not squared. This weighting is appropriate if you believe the data are drawn from a distribution that has relatively fat tails (leptokurtic; see Chapter 3) compared to the normal distribution. Other measures can be used that give more or less weight to large deviations. However, once we abandon RSS, we can no longer use the simple formulas to obtain estimates of the regression parameters. Instead, iterative computer techniques are necessary to find the combination of parameters that will minimize the residual (see Footnote 5 in Chapter 4). To illustrate robust regression, Figure 9.9 shows the Galápagos data with an additional outlier point. A standard linear regression for this new dataset gives a slope of 0.233, 30% shallower than the actual slope of 0.331 shown in Figure 9.4. Two different robust regression techniques, regression using leasttrimmed squares (Davies 1993) and regression using M-estimators (Huber 1981), were applied to the data shown in Figure 9.9. As in standard linear regression, these robust regression methods assume the X variable is measured without error. Least-trimmed squares regression minimizes the residual sums of squares (see Equation 9.5) by trimming off some percentage of the extreme observations. For a 10% trimming—removing the largest decile of the residual sums of squares—the predicted slope of the species-area relationship for the data illustrated in Figure 9.9 is 0.283, a 17% improvement over the basic linear regression. The intercept for the least-trimmed squares regression is 1.432, somewhat higher than the actual intercept. M-estimators minimize the residual: n ⎛ Y − X ib ⎞ residual = ∑ ρ ⎜ i + n log s s ⎟⎠ i =1 ⎝

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Figure 9.9 Three regressions for the species–area relationship of plant species on the Galápagos islands (see Table 8.2; see also Figures 8.6 and 9.3) with an added artificial outlier point. The standard linear regression (LS) is relatively sensitive to outliers. Whereas the original slope estimate was 0.331, the new slope is only 0.233, and has been dragged down by the outlier point. The least-trimmed squares (LTS) method discards the extreme 10% of the data (5% from the top and 5% from the bottom). The slope for the LTS regression is 0.283—somewhat closer to the original estimate of 0.331, although the intercept estimate is inflated to 1.432. The M-estimate weights the regression by the size of the residual, so that large outliers contribute less to the slope and intercept calculations. The M-estimator recovers the “correct” estimates for the slope (0.331) and the intercept (1.319), although the estimated variance is about double that of the linear regression that did not include the outlier. These robust regression methods are useful for fitting regression equations to highly variable data that include outliers.

where Xi and Yi are values for predictor and response variables, respectively; ρ = –log f, where f is a probability density function of the residuals ε that is scaled by a weight s, [f(ε/s)]/s, and b is an estimator of the slope β1. The value for b that minimizes Equation 9.34 for a given value of s is a robust M-estimator of the slope of the regression line β1. The M-estimator effectively weights the data points by their residuals, so that data points with large residuals contribute less to the slope estimate. It is, of course, necessary to provide an estimate of s to solve this equation. Venables and Ripley (2002) suggest that s can be estimated by solving n

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where ψ is the derivative of ρ used in Equation 9.34. Fortunately, all these computations are handled easily in software such as S-Plus or R that have built-in robust regression routines (Venables and Ripley 2002).

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M-estimate robust regression yields the correct slope of 0.331, but the variance around this estimate equals 0.25—more than three times as large as the estimate of the variance of the data lacking the outlier (0.07). The intercept is similarly correct at 1.319, but its variance is nearly twice as large (0.71 versus 0.43). Lastly, the overall estimate of the variance of the regression error equals 0.125, just over 30% larger than that of the simple linear regression on the original data. A sensible compromise strategy is to use the ordinary least-squares methods for testing hypotheses about the statistical significance of the slope and intercept. With outliers in the data, such tests will be relatively conservative because the outliers will inflate the variance estimate. However, robust regression could then be used to construct prediction intervals. The only caveat is that if the errors are actually drawn from a normal distribution with a large variance, the robust regression is going to underestimate the variance, and you may be in for an occasional surprise if you make predictions from the robust regression model. Quantile Regression

Simple linear regression fits a line through the center of a cloud of points, and is appropriate for describing a direct cause-and-effect relationship between the X and the Y variables. However, if the X variable acts as a limiting factor, it may impose a ceiling on the upper value of the Y variable. Thus, the X variable may control the maximum value of Y, but have no effect below this maximum. The result would be a triangle-shaped graph. For example, Figure 9.10 depicts the relationship between annual acorn biomass and an “acorn suitability index” measured for 43 0.2-hectare sample plots in Missouri (Schroeder and Vangilder 1997). The data suggest that low suitability constrains annual acorn biomass, but at high suitability, acorn biomass in a plot may be high or low, depending on other limiting factors. A quantile regression minimizes deviations from the fitted regression line, but the minimization function is asymmetric: positive and negative deviations are weighted differently: n

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As in robust regression, the function minimizes the absolute value of the deviations, so it isn’t as sensitive to outliers. However, the key feature is the multiplier h. The value of h is the quantile that is being estimated. If the deviation inside the absolute value sign is positive, it is multiplied by h. If the deviation is negative, it is multiplied by (1.0 – h). This asymmetric minimization fits a regression line through the upper regions of the data for large h, and through the lower regions for small h. If h = 0.50, the regression line passes through the center of the cloud of data and is equivalent to a robust regression using Equation 9.5.

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Figure 9.10 Illustration of quantile regression. The X variable is a measure of the acorn suitability of a forest plot, based on oak forest characteristics for n = 43 0.2-ha sample plots in Missouri (data from Schroeder and Vangilder 1997). The Y variable is the annual acorn biomass in the plot. Solid lines correspond to quantile regression slopes. The dashed line (= 50th percentile) is the standard least-squares regression that passes through the center of the cloud of points. Quantile regression is appropriate when there is a limiting factor that sets an upper ceiling on a response variable. (After Cade et al. 1999.)

Max 250

200 Annual acorn biomass (kg/ha)

272

150 95th 90th 100

75th 50th

25th 10th

0 0.0

0.2

0.4 0.6 Acorn suitability index

0.8

1.0

The result is a family of regression lines that characterize the upper and lower boundaries of the data set (Figure 9.10). If the X variable is the only factor affecting the Y variable, these quantile regressions will be roughly parallel to one another. However, if other variables come into play, the slopes of the regressions for the upper quantiles will be much steeper than for the standard regression line. This pattern would be indicative of an upper boundary or a limiting factor at work (Cade and Noon 2003). Some cautions are necessary with quantile regression. The first is that the choice of which quantile to use is rather arbitrary. Moreover, the more extreme the quantile, the smaller the sample size, which limits the power of the test. Also, quantile regressions will be dominated by outliers, even though Equation 9.36 minimizes their effect relative to the least-squares solution. Indeed, quantile regression, by its very definition, is a regression line that passes through extreme data points, so you need to make very sure these extreme values do not merely represent errors. Finally, quantile regression may not be necessary unless the hypothesis is really one of a ceiling or limiting factor. Quantile regression is often used with “triangular” plots of data. However, these data triangles may simply reflect heteroscedascity—a variance that increases at higher values of the X variable.

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Data transformations and careful analysis of residuals may reveal that a standard linear regression is more appropriate. See Thompson et al. (1996), Garvey et al. (1998), Scharf et al. (1998), and Cade et al. (1999) for other regression methods for bivariate ecological data. Logistic Regression

Logistic regression is a special form of regression in which the Y variable is categorical, rather than continuous. The simplest case is of a dichotomous Y variable. For example, we conducted timed censuses at 42 randomly-chosen leaves of the cobra lily (Darlingtonia californica), a carnivorous pitcher plant that captures insect prey (see Footnote 11 in Chapter 8). We recorded wasp visits at 10 of the 42 leaves (Figure 9.11). We can use logistic regression to test the hypothesis that visitation probability is related to leaf height.14 You might be tempted to force a regression line through these data, but even if the data were perfectly ordered, the relationship would not be linear. Instead, the best fitting curve is S-shaped or logistic, rising from some minimum value to a maximum asymptote. This kind of curve can be described by a function with two parameters, β0 and β1: p=

e β 0 + β1 X 1 + e β 0 + β1 X

(9.37)

The parameter β0 is a type of intercept because it determines the probability of success (Yi = 1) p when X = 0. If β0 = 0, then p = 0.5. The parameter β1 is similar to a slope parameter because it determines how steeply the curve rises to the maximum value of p = 1.0. Together, β0 and β1 specify the range of the X variable over which most of the rise occurs and determine how quickly the probability value rises from 0.0 to 1.0. The reason for using Equation 9.37 is that, with a little bit of algebra, we can transform it as follows: ⎛ p ⎞ ln ⎜ ⎟ = β 0 + β1 X ⎝1− p ⎠

(9.38)

14 Of course, we could use a much simpler t-test or ANOVA (see Chapter 10) to compare the heights of visited versus unvisited plants. However, an ANOVA for these data subtly turns cause and effect on its head: the hypothesis is that plant height influences capture probability, which is what is being tested with logistic regression. The ANOVA layout implies that the categorical variable visitation somehow causes or is responsible for variation in the continuous response variable leaf height.

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1.0 0.8 P (visitation)

274

0.6 0.4 0.2 0.0 0

40 Leaf height (cm)

Figure 9.11 Relationship between leaf height and wasp visitation for the carnivorous plant Darlingtonia californica. Each point represents a different plant in a population in the Siskiyou Mountains of southern Oregon (Ellison and Gotelli, unpublished data). The x-axis is the height of the leaf, a continuous predictor variable. The y-axis is the probability of wasp visitation. Although this is a continuous variable, the actual data are discrete, because a plant is either visited (1) or not (0). Logistic regression fits an S-shaped (= logistic) curve to these data. Logistic regression is used here because the response variable is discrete, so the relationship has an upper and lower asymptote. The model is fit using the logit transformation (Equation 9.38). The best-fit parameters (using maximum likelihood by iterative fitting) are βˆ 0 = –7.293 and βˆ 1 = 0.115. The P-value for a test of the null hypothesis that β1 = 0 is 0.002, suggesting that the probability of wasp visitation increases with increasing leaf height.

This transformation of the Y variable, called the logit transformation, converts the S-shaped logistic curve into a straight line. Although this transformation is indeed linear for the X variable, we cannot apply it directly to our data. If the data consist only of 0’s and 1’s for plants of different sizes, Equation 9.38 cannot be solved because ln[p/(1 – p)] is undefined for p = 1 or p = 0. But even if the data consist of estimates of p based on multiple observations of plants of the same size, it still would not be appropriate to use the least-squares estimators because the error term follows a binomial distribution rather than a normal distribution. Instead, we use a maximum likelihood approach. The maximum likelihood solution gives parameter estimates that make the observed values in the data set most probable (see Footnote 13 in Chapter 5). The maximum likelihood solution includes an estimate of the regression error variance, which can be used to test null hypotheses about parameter values and construct confidence intervals, just as in standard regression.

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For the Darlingtonia data the maximum likelihood estimates for the parameters are βˆ 0 = –7.293 and βˆ 1 = 0.115. The test of the null hypothesis that β1 = 0 generates a P-value of 0.002, suggesting that the probability of wasp visitation increases with leaf size. Non-Linear Regression

Although the least squares method describes linear relationships between variables, it can be adapted for non-linear functions. For example, the double logarithmic transformation converts a power function for X and Y variables (Y = aXb) into a linear relationship between log(X) and log(Y): log(Y) = log(a) + b × log(X). However, not all functions can be transformed this way. For example, many non-linear functions have been proposed to describe the functional response of predators—that is the change in predator feeding rate as a function of prey density. If a predator forages randomly on a prey resource that is depleted through time, the relationship between the number eaten (Ne) and the initial number (N0) is (Rogers 1972):

(

N e = N 0 1 − e a(Th N e −T )

)

(9.39)

In this equation, there are three parameters to be estimated: a, the attack rate, Th, the handling time per prey item, and T, the total prey handling time. The Y variable is Ne , the number eaten, and the X variable is N0 , the initial number of prey provided. There is no algebraic transformation that will linearize Equation 9.39, so the least-squares method of regression cannot be used. Instead, non-linear regression is used to fit the model parameters in the untransformed function. As with logistic regression (a particular kind of non-linear regression), iterative methods are used to generate parameters that minimize the least-squares deviations and allow for hypothesis tests and confidence intervals. Even when a transformation can produce a linear model, the least-squares analysis assumes that the error terms εi are normally distributed for the transformed data. But if the error terms are normally distributed for the original function, the transformation will not preserve normality. In such cases, non-linear regression is also necessary. Trexler and Travis (1993) and Juliano (2001) provide good introductions to ecological analyses using non-linear regression. Multiple Regression

The linear regression model with a single predictor variable X can easily be extended to two or more predictor variables, or to higher-order polynomials of a single predictor variable. For example, suppose we suspected that species rich-

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ness peaks at intermediate island sizes, perhaps because of gradients in disturbance frequency or intensity. We could fit a second-order polynomial that includes a squared term for island area: Yi = β 0 + β1 X i + β 2 X i2 + ε i

(9.40)

This equation describes a function with a peak of species richness at an intermediate island size.15 Equation 9.40 is an example of a multiple regression, because there are now actually two predictor variables, X and X 2, which contribute to variation in the Y variable. However, it is still considered a linear regression model (albeit a multiple linear regression) because the parameters βi in Equation 9.40 are solvable using linear equations. If the data are modeled as a simple linear function, we ignore the polynomial term, and the systematic component of variation from X 2 is incorrectly pooled with the error term: ε i′ = β 2 X i2 + ε i

(9.41)

A more familiar example of multiple regression is when two or more distinct predictor variables are measured for each replicate. For example, in a study of variation in species richness of ants (S) in New England bogs and forests (Gotelli and Ellison 2002a,b), we measured the latitude and elevation of each study site and entered both variables in a multiple regression equation: log10 ( forest S) = 4.879 − 0.089(latitude) − 0.001(elevation)

(9.42)

Both slope parameters are negative because species richness declines at higher elevations and higher latitudes. The parameters in this model are called partial regression parameters because the residual sums of squares from the other variables in the model have already been accounted for statistically. For example, the parameter for latitude could be found by first regressing species richness on elevation, and then regressing the residuals on latitude. Conversely, the parameter for elevation could be found by regressing the residuals from the richness-latitude regression on elevation.

The peak can be found by taking the derivative of Equation 9.40 and setting it equal to zero. Thus, the maximum species richness would occur at X = β1/β2. Although this equation produces a non-linear curve for the graph of Y versus X, the model is still a linear sum of the form Σβi Xi , where Xi is the measured predictor variable (which itself may be a transformed variable) and βi ’s are the fitted regression parameters.

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Because these partial regression parameters are based on residual variation not accounted for by other variables, they often will not be equivalent to parameters estimated in simple regression models. For example, if we regress species richness on latitude only, the result is log10 ( forest S) = 5.447 − 0.105(latitude)

(9.43)

And, if we regress species richness on elevation only, we have log10 ( forest S) = 1.087 − 0.001(elevation)

(9.44)

With the exception of the elevation term, all of the parameters differ among the simple linear regressions (Equations 9.43 and 9.44) and the multiple regression (Equation 9.42). In linear regression with a single predictor variable, the function can be graphed as a line in two dimensions (Figure 9.12). With two predictor variables, the multiple regression equation can be graphed as a plane in a three-dimensional coordinate space (Figure 9.13). The “floor” of the space represents the two axes for the two predictor variables, and the vertical dimension represents the response variable. Each replicate consists of three measurements (Y variable, X1 variable, X2 variable), so the data can be plotted as a cloud of points in the three

Forest ant species number

(A)

(B) 100

100

1 41

42 43 44 Degrees N latitude

100 200 300 400 500 600 Elevation (m)

Figure 9.12 Forest ant species density as a function of (A) latitude and (B) elevation. Each point (n = 22) is the number of ant species recorded in a 64-m2 forest plot in New England (Vermont, Massachusetts, and Connecticut). The least-squares regression line is shown for each variable. Note the log10 transformation of the y-axis. For the latitude regression, the equation is log10(ant species number) = 5.447 – 0.105 × (latitude); r 2 = 0.334. For the elevation regression, the equation is log10(ant species number) = 1.087 – 0.001 × (elevation); r 2 = 0.353. The negative slopes indicate that ant species richness declines at higher latitudes and higher elevations. (Data and sampling details in Gotelli and Ellison 2002a,b.)

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2.0 log10(Forest ant species richness)

278

1.6 1.2 0.8 600 500 400 El ev 300 ati on 200 100 (m ) 0

46 45 44 43 42 41

ees Degr

itude

N lat

Figure 9.13 Three-dimensional representation of multiple regression data. The least squares solution is a plane that passes through the cloud of data. Each point (n = 22) is the number of ant species recorded in a 64-m2 forest plot in northern New England (Vermont, Massachusetts, and Connecticut). The X variable is the plot latitude (the first predictor variable), the Y variable is the plot elevation (the second predictor variable), and the Z variable is the log10 of the number of ant species recorded (the response variable). A multiple regression equation has been fit to the equation: log10(ant species number) = 4.879 – 0.089 × (latitude) – 0.001 × (elevation); r 2 = 0.583. Notice that the r 2 value, which is a measure of the fit of the data to the model, is larger for the multiple regression model than it is for either of the simple regression models based on each predictor variable by itself (see Figure 9.12). The solution to a linear regression with one predictor variable is a straight line, whereas the solution to a multiple regression with two predictor variables is a plane, which is shown in this three-dimensional rendering. Residuals are calculated as the vertical distance from each datum to the predicted plane. (Data and sampling details in Gotelli and Ellison 2002a,b.)

dimensional space. The least-squares solution passes a plane through the cloud of points. The plane is positioned so that the sum of the squared vertical deviations of all of the points from the plane is minimized. The matrix solutions for multiple regression are described in the Appendix, and their output is similar to that of simple linear regression: least-squares parameter estimates, a total r 2, an F-ratio to test the significance of the entire model, and error variances, confidence intervals, and hypothesis tests for each of the coefficients. Residual analysis and tests for outliers and influential points can be carried out as for single linear regression. Bayesian or Monte Carlo methods also can be used for multiple regression models. However, a new problem arises in evaluating multiple regression models that was not a problem in simple linear regression: there may be correlations among the predictor variables themselves, referred to as multicollinearity (see Graham 2003). Ideally, the predictor variables are orthogonal to one another: all values

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of one predictor variable are found in combination with all values of the second predictor variable. When we discussed the design of two-way experiments in Chapter 7, we emphasized the importance of ensuring that all possible treatment combinations are represented in a fully crossed design. The same principle holds in multiple regression: ideally, the predictor variables should not themselves be correlated with one another. Correlations among the predictor variables make it difficult to tease apart the unique contributions of each variable to the response variable. Mathematically, the least-squares estimates also start to become unstable and difficult to calculate if there is too much multicollinearity between the predictor variables. Whenever possible, design your study to avoid correlations between the predictor variables. However, in an observational study, you may not be able to break the covariation of your predictor variables, and you will have to accept a certain level of multicollinearity. Careful analysis of residuals and diagnostics is one strategy for dealing with covariation among predictor variables. Another strategy is to mathematically combine a set of intercorrelated predictor variables into a smaller number of orthogonal variables with multivariate methods such as principal components or discriminant analysis (see Chapter 12). Is multicollinearity a problem in the ant study? Not really. There is very little correlation between elevation and latitude, the two predictor variables for ant species richness (Figure 9.14). Path Analysis

All of the regression models that we have discussed so far (linear regression, robust regression, quantile regression, non-linear regression, and multiple regression) begin with the designation of a single response variable and one or more predictor variables that may account for variation in the response. But in reality, many models of ecological processes do not organize variables into a single

600

Elevation (m)

500 400 300 200 100 0 41

43 44 Degrees N latitude

Figure 9.14 Lack of collinearity among the predictor variables strengthens multiple regression analyses. Each point represents the latitude and elevation in a set of 22 forest plots for which ant species richness had been measured (see Figure 9.12). Although elevation and latitude can be used simultaneously as predictors in a multiple regression model (see Figure 9.13), the fit of the model can be compromised if there are strong correlations among the predictor variables (collinearity). In this case, there is only a very weak correlation between the predictor variables latitude and elevation (r 2 = 0.032; F1,20 = 0.66, P = 0.426). It is always a good idea to test for correlations among predictor variables when using multiple regression. (Data and sampling details in Gotelli and Ellison 2002a,b.)

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Figure 9.15 Path analysis for passive colonization and food web models of Sarracenia

inquilines. Each oval represents a different taxon that can be found in the leaves of the northern pitcher plant, Sarracenia purpurea. Captured insect prey forms the basis for this complex food web, which includes several trophic levels. The underlying data consist of abundances of organisms measured in leaves that had different water levels in each leaf (Gotelli and Ellison 2006). Treatments were applied for one field season (May–August 2000) in a press experiment (see Chapter 6) to 50 plants (n = 118 leaves). The abundance data were fit to two different path models, which represent two different models of community organization. (A) In the passive colonization model, abundances of each taxon depend on the volume of water in each leaf and the level of prey in the leaf, but no interactions among taxa are invoked. (B) In the trophic model, the abundances are determined entirely by trophic interactions among the inquilines. The number associated with each arrow is the standardized path coefficient. Positive coefficients indicate that increased prey abundance leads to increased predator abundance. Negative coefficients indicate that increased predator abundance leads to decreased prey abundance. The thickness of each arrow is proportional to the size of the coefficient. Positive coefficients are blue lines, negative coefficients are gray lines.

response variable and multiple predictor variables. Instead, measured variables may act simultaneously on each other in cause-and-effect relationships. Rather than isolate the variation in a single variable, we should try to explain the overall pattern of covariation in a set of continuous variables. This is the goal of path analysis. Path analysis forces the user to specify a path diagram that illustrates the hypothesized relationships among the variables. Variables are connected to one another by single- or double-headed arrows. Variables that do not interact directly are not connected by arrows. A path diagram of this sort represents a mechanistic hypothesis of interactions in a system of variables.16 It also represents a statistical hypothesis about the structure of the variance-covariance matrix of these variables (see Footnote 5 in this chapter). Partial regression parameters can then be estimated for the individual paths in the diagram, and summary goodness-of-fit statistics can be derived for the overall evaluation of the model. For example, path analysis can be used to test different models for community structure of the invertebrate species that co-occur in pitcher-plant leaves (Gotelli and Ellison 2006). The data consist of repeated censuses of entire communities that live in pitcher plant leaves (n = 118 leaves). The replicates are the 16

Sewall Wright

Path analysis was first introduced by the population geneticist Sewall Wright (1889–1988) as a method for analyzing genetic covariation and patterns of trait inheritance. It has long been used in the social sciences, but has become popular with ecologists only recently. It has some of the same strengths and weaknesses as multiple regression analysis. See Kingsolver and Schemske (1991), Mitchell (1992), Petraitis et al. (1996), and Shipley (1997) for good discussions of path analysis in ecology and evolution.

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Passive colonization model

(A) Fletcherimyia (flesh fly)

–0.010

Wyeomyia (mosquito)

0.149

Volume

0.433

–0.068

Habrotrocha (rotifer)

0.018

–0.039

–0.089

Prey

0.366

–0.039

Metriocnemus (midge)

0.040

–0.002

0.104

Sarraceniopus (mite)

(B)

0.162

Protozoa

Food web model Fletcherimyia (flesh fly) 0.271 Wyeomyia (mosquito)

–0.047 Habrotrocha (rotifer)

–0.252 –0.063

0.010 Protozoa

Metriocnemus (midge)

Sarracenioppus (mite)

0.177 0.094

0.155 Prey

individual leaves that were censused, and the continuous variables are the average abundance of each invertebrate species. One model to account for community structure is a passive colonization model (Figure 9.15A). In this model, the abundance of each species is determined large-

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ly by the volume of the pitcher-plant leaf and the food resources (ant prey) available in the leaf. This model does not include any explicit interactions between the resident species, although they do share the same food resources. A second model to account for community structure is a food web model, in which trophic interactions between species are important in regulating their abundances (Figure 9.15B). Prey resources occur at the bottom of the trophic chain, and these are processed by mites, midges, protozoa, and flesh flies. Although flesh flies are the top predator, some of the species in this community have multiple trophic roles. The passive colonization model and the food web model represent two a priori hypotheses about the forces controlling inquiline abundances. Figure 9.15 illustrates the path coefficients associated with each model. In the passive colonization model, the largest coefficients are positive ones between leaf volume and midge abundance, leaf volume and prey abundance, and prey abundance and protozoa abundance. The confidence intervals of most coefficients in the model bracket zero. In the food web model, there are large positive coefficients between prey and midges, between prey and protozoa, and between mosquito abundance and fleshfly abundance. A strong negative coefficient links mosquitoes and their rotifer prey. As in the passive colonization model, confidence intervals for many of the coefficients bracket zero. Path analysis is very much a Bayesian approach to modeling and can be implemented in a full Bayesian framework, in which path coefficients are treated as random variables (Congdon 2002). It requires the user to specify a priori hypotheses in the form of path diagrams with arrows of cause and effect. In contrast, simple and multiple regression is more frequentist in spirit: we propose a parsimonious linear model for the data and test the simple null hypothesis that the model coefficients (βi) do not differ from zero.

Model Selection Criteria Path analysis and multiple regression present a problem that has not arisen so far in our survey of statistical methods: how to choose among alternative models. For both multiple regression and path analysis, coefficients and statistical significance can be calculated for any single model. But how do we choose among different candidate models? For multiple regression, there may be many possible predictor variables we could use. For example, in the forest ant study, we also measured vegetation structure and canopy cover that could potentially affect ant species richness. Should we just use all of these variables, or is there some way to select a subset of predictor variables to most parsimoniously account for variation in the response variable? For path analysis, how do we decide which of two or more a priori models best fits the data?

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Model Selection Methods for Multiple Regression

For a dataset that has one response variable and n predictor variables, there are (2n – 1) possible regression models that can be created. These range from simple linear models with only a single predictor variable to a fully saturated multiple regression model that has all n predictor variables present.17 It would seem that the best model would be the one that has the highest r 2 and explains most of the variation in the data. However, you would find that the fully saturated model always has the highest r 2. Adding variables to a regression model never increases the RSS and usually will decrease it, although the decrease can be pretty small when adding certain variables to the model. Why not use only the variables for which the slope coefficients (βi) in the saturated model are statistically different from zero? The problem here is that coefficients—and their statistical significance—depend on which other variables are included in the model. There is no guarantee that the reduced model containing only the significant regression coefficients is necessarily the best model. In fact, some of the coefficients in the reduced model may no longer be statistically significant, especially if there is multicollinearity among the variables. Variable selection strategies include forward selection, backward elimination, and stepwise methods. In forward selection, we begin adding variables one at a time, and stop at a certain criterion. In backward elimination models, we start with the fully saturated model (all variables included) and begin eliminating variables one at a time. Stepwise models include both forward and backward steps to make comparisons by swapping variables in and out of the model and evaluating changes in the criterion. Typically, two criteria are used to decide when to stop adding (or removing) variables. The first criterion is a change in the F-ratio of the fitted model. The F-ratio is a better choice than r 2 or the RSS because the change in the F-ratio depends both on the reduction in r 2 and on the number of parameters that are included in the model. In forward selection, we continue adding variables until

There are actually even more possible models and variables than this. For example, if we have two predictor variables X1 and X2, we can create a composite variable X1X2. The regression model Yi = β0 +β1X1i + β2X2i + β3X1i X2i + εi has coefficients for both the main effects of X1 and X2, and a coefficient for the interaction between X1 and X2. This term is analogous to the interaction between discrete variables in a two-way ANOVA (see Chapter 10). As we have noted before, statistical software will cheerfully accept any linear model you care to construct; it is your responsibility to examine the residuals and consider the biological implications of different model structures.

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the increase in the F-ratio falls below a specified threshold. In backwards elimination, we eliminate variables until there is too large a drop in the F-ratio. The second criterion for variable selection is tolerance. Variables are removed from a model or not added in if they create too much multicollinearity among the set of predictor variables. Typically, the software algorithm creates a multiple regression in which the candidate variable is the response variable, and the other variables already in the model are the predictor variables. The quantity (1 – r 2) in this context is the tolerance. If the tolerance is too low, the new variable shows too much correlation with the existing set of predictor variables and is not included in the equation. All computer packages contain default cutoff points for F-ratios and tolerances, and they will all generate a reasonable set of predictor variables in multiple regression. However, there are two problems with this approach. The first problem is that there is no theoretical reason why variable selection methods will necessarily find the best subset of predictor variables in multiple regression. The second problem is that these selection methods are based on optimization criteria, so they leave you with only a single best model. The algorithms don’t give you alternative models that may have had very similar F-ratios and regression statistics. These alternative models include different variables, but are statistically indistinguishable from the single best-fitting model. If the number of candidate variables is not too large (< 8), a good strategy is to calculate all possible regression models and then evaluate them on the basis of F-ratios or other least-squares criteria. This way, you can at least see if there is a large set of similar models that may be statistically equivalent. You may also find patterns in the variables—perhaps one or two variables always show up in the final set of models—that can help you choose a final subset. Our point here is that you cannot rely on automated or computerized statistical analyses to sift through a set of correlated variables and identify the correct model for your data. The existing selection methods are reasonable, but they are arbitrary. Model Selection Methods in Path Analysis

Path analysis forces the investigator to propose specific models that are then evaluated and compared. The selection of the “best” model in path analysis presumes that the “correct” model is included among the alternatives you are evaluating. If none of the proposed path models is correct, you simply are choosing an incorrect model that fits relatively well compared to the alternative incorrect models. In both path analysis and multiple regression, the problem is to choose a model that has enough variables to properly account for the variation in the data and minimize the residual sum of squares, but not so many variables that you

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Model Selection Criteria

are fitting equations to random noise in the data. Information criterion statistics balance a reduction in the sum of squares with the addition of parameters to the model. These methods penalize models with more parameters in them, even though such models inevitably reduce the sum of squares. For example, an information criterion index for multiple regression is the adjusted r 2:

(

⎛ n −1 ⎞ 2 2 radj = 1− ⎜ ⎟ 1− r ⎝ n − p⎠

)

(9.45)

where n is the sample size and p is the number of parameters in the model (p = 2 for simple linear regression with a slope parameter and an intercept parameter). Like r 2, this number increases as more of the residual variation is 2 explained. However, radj also decreases as more parameters are added to the 2 model. Therefore, the model with the largest radj is not necessarily the model with the most parameters. In fact, if the model has lots of parameters and fits the data 2 poorly, radj can even become negative. For path analysis, the Akaike information criterion, or AIC, can be calculated as AIC = −2 log ⎡⎣ L(θˆ | data)⎤⎦ + 2 K

(9.46)

where L(θˆ⎥ y) is the likelihood of the estimated model parameter (θˆ ) given the data, and K is the number of parameters in the model. In path analysis, the number of parameters in the model is not simply the number of arrows in the path diagram. The variances of each variable have to be estimated as well. The AIC can be thought of as a “badness-of-fit” measure, because the larger the number, the more poorly the data fit the variance-covariance structure that is implied by the path diagram. For example, the random colonization model for the inquilines has 21 parameters and a cross-validation index (an AIC measure) of 0.702, whereas the food web model has 15 parameters and a crossvalidation index of 0.466. Bayesian Model Selection

Bayesian analysis can be used to compare different hypotheses or models (such as the regression models with and without the slope term β1; see Equations 9.20 and 9.21). Two methods have been developed (Kass and Raftery 1995; Congdon 2002). The first, called Bayes’ factors, estimates the relative likelihood of one hypothesis or model relative to another hypothesis or model. As in a full Bayesian analysis, we have prior probabilities P(H0) and P(H1) for our two hypothesis or models. From Bayes’ Theorem (see Chapter 1), the posterior probabilities

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(9.47)

We define the prior odds ratio, or the relative probability of one hypothesis versus the other hypothesis, as P(H0)/P(H1)

(9.48)

If these are the only two alternatives, then P(H0) + P(H1) = 1 (by the First Axiom of Probability; see Chapter 1). Equation 9.48 expresses how much more or less likely one hypothesis is relative to the other before we conduct our experiment. At the start of the study, we expect the prior probabilities of each hypothesis or model to be roughly equal (otherwise, why waste our time gathering the data?), and so Equation 9.48 ≈ 1. After the data are collected, we calculate posterior probabilities. The Bayes factor is the posterior odds ratio: P(H0 | data) / P(H1 | data)

(9.49)

If Equation 9.49 >>1, then we would have reason to believe that H0 is favored over H1, whereas if Equation 9.49 C = U; Figure 10.2B). This result suggests that the treatment has a significant effect on the response variable, and that the result does not represent an artifact of the design. Note that the treatment effect would have been inferred erroneously if we had (incorrectly) set the experiment up without the controls and only compared the treatment and unmanipulated groups. Alternatively, suppose that the treatment and control groups have similar means, but they are both elevated above the unmanipulated plots (T = C > U; Figure 10.2C). This pattern suggests that the treatment effect is not important and that the difference among the means reflects a handling effect or other artifact of the manipulation. Finally, suppose that the three means differ from one another, with the treatment group mean being the highest and the unmanipulated group mean being the lowest (T > C > U; Figure 10.2D). In this case, there is evidence of a handling effect because C > U. However, the fact that T > C means that the treatment effect is also real and does not represent just handling artifacts. Once you have established the pattern in your data using ANOVA and the graphical output, the next step is to determine whether that pattern tends to support or refute the scientific hypothesis you are evaluating (see Chapter 4). Always keep in mind the difference between statistical significance and biological significance!

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After ANOVA: Plotting and Understanding Interaction Terms

(A)

(B)

Response variable

20 18 16 14 12 10 (C)

(D)

Response variable

20 18 16 14 12 10 Unmanipulated Control

Treatment

Unmanipulated Control

Treatment

Figure 10.2 Possible outcomes of a hypothetical experiment with three treatments and a one-way ANOVA layout. Unmanipulated plots are not altered in any way except for effects that occur during sampling. Control plots do not receive the treatment, but may receive a sham treatment to mimic handling effects. The treatment group has the manipulation of interest, but also potentially includes handling effects. The height of each bar represents the mean response for the group, and the vertical line indicates 1 standard deviation around the mean. (A) The ANOVA test for treatment effects is non-significant, and the differences among groups in the mean response can be attributed to pure error. (B–D) The ANOVA test is always significant, but the pattern—and hence the interpretation—differs in each case. (B) The treatment mean is elevated compared to the control and unmanipulated replicates, indicating a true treatment effect. (C) Both the control and the treatment group means are elevated relative to the unmanipulated plot, indicating a handling effect, but no distinct treatment effect. (D) There is evidence of a handling effect because the mean of the control group exceeds that of the unmanipulated plot, even though the desired (biological) treatment was not applied. However, above and beyond this handling effect, there appears to be a treatment effect because the mean of the treatment group is elevated relative to the mean of the control group. A priori contrasts or a posteriori comparisons can be used to pinpoint which groups are distinctly different from another. Our point here is that the results of the ANOVA might be identical, but the interpretation depends on the particular pattern of the group means. ANOVA results cannot be interpreted meaningfully without a careful examination of the pattern of means and variances of the treatment groups.

Plotting Results from Two-Way ANOVAs

For plotting the results of two-way ANOVAs, simple bar graphs also can be used, but they are harder to interpret graphically and do not display the main effects and interaction effects very well. We suggest the following protocol for plotting the means in a two-way ANOVA:

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1. Establish a plot in which the y-axis is the continuous response variable, and the x-axis represents the different levels for the first factor in the experiment. 2. To represent the second factor in the experiment, use a different symbol or color for each treatment level. Each symbol, placed over the appropriate category label on the x-axis, represents the mean of a particular treatment combination. There will be a total of a × b symbols where a is the number of treatments in the first factor and b is the number of treatments in the second factor. 3. Align each symbol above the appropriate factor label to establish the combination of treatments represented. 4. Use a (colored) line to connect the symbols across the levels of the first factor. 5. To plot standard deviations use vertical lines pointing upward for the higher treatment levels and vertical lines pointing downward for lower treatment levels. If the figure becomes too cluttered, error bars can be illustrated for only some of the data series. Figure 10.3 shows just such a plot for the hypothetical two-way barnacle experiment we described. The x-axis gives the four levels of the predation treatment

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Figure 10.3 Possible outcomes for a hypothetical experiment testing for the effects of substrate and predation treatment on barnacle recruitment. Each symbol represents a different treatment combination mean. Predation treatments are indicated by the x-axis label and substrate treatments are indicated by the different colors. Vertical bars indicate standard errors or standard deviations (which must be designated in the figure legend). Each panel represents the pattern associated with a particular statistical outcome. (A) Neither treatment effects nor the interaction are significant, and the means of all the treatment combinations are statistically indistinguishable. (B) The predation effect is significant, but the substrate effect and interaction effect are not. In this graph, treatment means are highest for the predator exclusion and lowest for the predator inclusion, with a similar pattern for all substrates. (C) The substrate effect is significant, but the predator effect and the interaction effect are not. There are no differences in the means of the predation treatment, but recruitment is always highest on the granite substrates and lowest on the cement substrates, regardless of the predation treatment. (D) Both predation and substrate effects are significant, but the interaction effect is not. Means depend on both the substrate and the predation treatment, but the effect is strictly additive, and the profiles of the means are parallel across substrates and treatments. (E) Significant interaction effect. Means of the treatments differ significantly, but there is no longer a simple additive effect of either predation or substrate. The ranking of the substrate means depends on the predation treatment, and the ranking of the predation means depends on the substrate. Main effects may not be significant in this case because treatment averages across substrates or predation treatments do not necessarily differ significantly. (F) The interaction effect is significant, which means that the effect of the predation treatment depends on the substrate treatment and vice versa. In spite of this interaction, it is still possible to talk about the general effects of substrate on recruitment. Regardless of the predation treatment, recruitment is always highest on granite substrates and lowest on cement substrates. The interaction effect is statistically significant, but the profiles for the means do not actually cross.

After ANOVA: Plotting and Understanding Interaction Terms

(A) Number of barnacle recruits

50 45

50 45 40

(C)

(D)

Predation: NS Substrate: P < 0.05 Predation × Substrate: NS

Predation: P < 0.05 Substrate: NS Predation × Substrate: NS

Predation: P < 0.05 Substrate: P < 0.05 Predation × Substrate: NS

35 20 30 10

25 20

(E) 50 Number of barnacle recruits

(B)

50 Number of barnacle recruits

Predation: NS Substrate: NS Predation × Substrate: NS

Cement Slate Granite

329

(F) Predation: NS Substrate: NS Predation × Substrate: P < 0.05

Predation: NS Substrate: P < 0.05 Predation × Substrate: P < 0.05

10 Unmanipulated Control

Inclusion

Exclusion

Unmanipulated Control Inclusion

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(unmanipulated, control, predator exclusion, predator inclusion). Three colors (blue, gray, black) are used to indicate the three levels of the substrate treatment (cement, slate, and granite). For each substrate type, the means for the four levels of the predation treatment are connected by a line. Once again, we should consider the different outcomes of the ANOVA and what the associated graphs would look like. In this case, there are several possibilities, because we now have two hypothesis tests for the main effects of predation and substrate and a third hypothesis test for the interaction between these two. As before, the simplest scenario is the one in which neither the two main effects nor the interaction term are statistically significant. If the sample sizes are reasonably large, this will also be the case that is the messiest to plot because all of the treatment group averages will be very similar to one another, and the points for the means may be closely superimposed (Figure 10.3A).

NO SIGNIFICANT EFFECTS

ONE SIGNIFICANT MAIN EFFECT Next, suppose that the main effect for predation is significant, but the substrate effect and interaction are not. Therefore, the means of each predation treatment, averaged over the three substrate treatments, are significantly different from one another. In contrast, the means of the substrate types, averaged over the different predation treatments are not substantially different from one another. The graph will show distinctly different clumps of treatment means at each level of predation treatment, but the means of each substrate type will be nearly identical at each predation level (Figure 10.3B). Conversely, suppose the substrate effect is significant, but the predation effect is not. Now the means of each of the three substrate treatments are significantly different, averaging over the four predation treatments. However, the averages for the predation treatments do not differ. The three connected lines for the substrate types will be well separated from one another, but the slopes of those lines will be basically flat because there is no effect of the four treatments (Figure 10.3C). TWO SIGNIFICANT MAIN EFFECTS The next possibility is a significant effect of predation and of substrate, but no interaction term. In this case, the profile of the mean responses is again different for each of the substrate types, but it is no longer flat across each of the predation treatments. A key feature of this graph is that the lines connecting the different treatment groups are parallel to one another. When the treatment profiles are parallel, the effects of the two factors are strictly additive: the particular treatment combination can be predicted

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After ANOVA: Plotting and Understanding Interaction Terms

knowing the average effects of each of the two individual treatments. Additivity of treatment effects and a parallel treatment profile are diagnostic for an ANOVA in which both of the main effects are significant and the interaction term is not significant (Figure 10.3D). The final possibility we will consider is that the interaction term is significant, but neither of the two main effects are. If the interactions are strong, the lines of the profile plot may cross one another (Figure 10.3E). If the interactions are weak, the lines may not cross one another, although they are no longer (statistically) parallel (Figure 10.3F).

SIGNIFICANT INTERACTION EFFECT

Understanding the Interaction Term

With a significant interaction term, the means of the treatment groups are significantly different from one another, but we can no longer describe a simple additive effect for either of the two factors in the design. Instead, the effect of the first factor (e.g., predation) depends on the level of the second factor (e.g., substrate type). Thus, the differences among the substrate types depend on which predation treatment is being considered. In control and unmanipulated plots, recruitment was highest on granite substrates, whereas in predator inclusion and exclusion plots, recruitment was highest on slate. Equivalently, we can say that the effect of the predator treatment depends on the substrate. For the granite substrate, abundance is highest in the controls, whereas for the slate substrate, abundance is highest in the predator inclusion and exclusion treatments. The graphic portrayal of the interaction term as non-parallel plots of treatment means also has an algebraic interpretation. From Table 10.6, the sum of squares for the interaction term in the two-way ANOVA is a

SS AB = ∑ ∑ ∑ (Yij − Yi − Y j + Y )2

(10.25)

i=1 j =1 k =1

– Equivalently, we can add and subtract a term for Y , giving a

[

]

SS AB = ∑ ∑ ∑ (Yij − Y ) − (Yi − Y ) − (Y j − Y ) i=1 j =1 k =1

(10.26)

– – The first term (Y ij – Y ) in this expanded expression represents the deviation of – – each treatment group mean from the grand average. The second term (Y i – Y ) represents the deviation from the additive effect of Factor A, and the third – – term (Y j – Y ) represents the additive effect of Factor B. If the additive effects of Factors A and B together account for all of the deviations of the treatment

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means from the grand mean, then the interaction effect is zero. Thus, the interaction term measures the extent to which the treatment means differ from the strictly additive effects of the two main factors. If there is no interaction term, then knowing the substrate type and knowing the predation treatment would allow us to predict perfectly the response when these factors are combined. But if there is a strong interaction, we can no longer predict the combined effect, even if we understand the behavior of the solitary factors.8 It is clear why the interaction term is significant in Figure 10.3E, but why should the main effects be non-significant in this case? The reason is that, if you average the means across each predation treatment or across each substrate type, they would be approximately equal, and there would not be consistent differences for each factor considered in isolation. For this reason, it is sometimes claimed that nothing can be said about main effects when interaction terms are significant. This statement is only true for very strong interactions, in which the profile curves cross one another. However, in many cases, there may be overall trends for single factors even when the interaction term is significant. For example, consider Figure 10.3F, which would certainly generate a statistically significant interaction term in a two-way ANOVA. The significant interaction tells us that the differences between the means for the substrate types depend on the predation treatment. However, in this case the interaction arises mostly because the cement substrate × predator exclusion treatment has a mean that is very small relative to all other treatments. In all of the predation treatments, the granite substrate (black line) has the highest recruitment and the cement substrate (blue line) has the lowest recruitment. In the control treatment the differences among the substrate means are relatively small, whereas in the predator exclusion treatment, the differences among the substrate means are relatively large. Again, we emphasize that ANOVA results cannot be interpreted properly without reference to the patterns of means and variances in the data. Finally, we note that data transformations may sometimes eliminate significant interaction terms. In particular, relationships that are multiplicative on a

A morbid example of a statistical interaction is the effect of alcohol and sedatives on human blood pressure. Suppose that alcohol lowers blood pressure by 20 points and sedatives lower blood pressure by 15 points. In a simple additive world, the combination of alcohol and sedatives should lower blood pressure by 35 points. Instead, the interaction of alcohol and sedatives can lower blood pressure by 50 points or more and is often lethal. This result could not have been predicted simply by understanding their effects separately. Interactions are a serious problem in both medicine and environmental science. Simple experimental studies may quantify the effects of single-factor environmental stressors such as elevated CO2 or increased temperature, but there may be strong interactions between these factors that can cause unexpected outcomes.

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After ANOVA: Plotting and Understanding Interaction Terms

linear scale are additive on a logarithmic scale (see discussions in Chapters 3 and 8), and the logarithmic transformation can often eliminate a significant interaction term. Certain ANOVA designs do not include interaction terms, and we have to assume strict additivity in these models. Plotting Results from ANCOVAs

We will conclude this section by considering the possible outcomes for an ANCOVA. The ANCOVA plot should use the continuous covariate variable plotted on the x-axis, and the Y variable plotted on the y-axis. Each point represents an independent replicate, and different symbols or colors should be used to indicate replicates in different treatments. Initially, fit a linear regression line to each treatment group for a general plot of the results. We will present a single example in which there is one covariate and three treatment levels being compared. Here are the possibilities: COVARIATE, TREATMENT, AND INTERACTION NON-SIGNIFICANT

In this case, the three regression lines do not differ from one another, and the slope of those lines is not different from zero. The fitted intercept of each regression effectively estimates the average of the Y values (Figure 10.4A).

COVARIATE SIGNIFICANT, TREATMENT AND INTERACTION NON-SIGNIFICANT

In this case, the three regression lines do not differ from one another, but now the slope of the common regression line is significantly different from zero. The results suggest that the covariate accounts for variation in the data, but there are no differences among treatments after the effect of the covariate is (statistically) removed (Figure 10.4B).

TREATMENT SIGNIFICANT, COVARIATE AND INTERACTION TERM NON-SIGNIFICANT In this case, the regression lines again have equivalent zero slopes, but now the intercepts for the three levels are different, indicating a significant treatment effect. Because the covariate does not account for much of the variation in the data, the result is qualitatively the same as if a one-way ANOVA were used and the covariate measures ignored (Figure 10.4C). TREATMENT AND COVARIATE SIGNIFICANT, INTERACTION TERM NON-SIGNIFICANT In this case, the regression lines have equivalent non-zero slopes and the intercepts are significantly different. The covariate does account for some of the variation in the data, but there is still residual variation that can be attributed to the treatment effect. This result is equivalent to fitting a single regression line to the entire data set, and then using a one-way ANOVA on the residuals to test for treatment effects. When this result is obtained, it is appropriate to fit a common regression

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Y (response variable)

(A)

(B)

Treatment: NS Covariate: NS Treatment × Covariate: NS

Y (response variable)

(C)

(D)

Treatment: P < 0.05 Covariate: NS Treatment × Covariate: NS

(E)

Y (response variable)

Treatment: NS Covariate: P < 0.05 Treatment × Covariate: NS

Treatment: P < 0.05 Covariate: P < 0.05 Treatment × Covariate: NS

(F) Treatment: NS Covariate: NS Treatment × Covariate: P < 0.05

X (covariate)

Treatment: P < 0.05 Covariate: NS Treatment × Covariate: P < 0.05

X (covariate)

slope and use Equation 10.18 to estimate the adjusted treatment means (Figure 10.4D). INTERACTION TERM SIGNIFICANT This case represents heterogeneous regression slopes, in which it is necessary to fit a separate regression line, with its own slope

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Comparing Means

Figure 10.4 Possible outcomes for experiments with ANCOVA designs. In each panel, different symbols indicate replicates in each of three treatment groups. Each panel indicates a different possible experimental outcome with the associated ANCOVA result. (A) No significant treatment or covariate effects. The data are best fit by a single grand average with sampling error. (B) Covariate effect significant, no significant treatment effect. The data are best fit by a single regression line with no differences among treatments. (C) Treatment effect significant, no significant covariate. The covariate term is not significant (regression slope = 0), and the data are best fit by a model with different group means, as in a simple one-way ANOVA. (D) Treatment and covariate are both significant. The data are best fit by a regression model with a common slope, but different intercepts for each treatment group. This model is used to calculate adjusted means, which are estimated for the grand mean of the covariate. (E) A treatment × covariate interaction in which the order of the treatment means differs for different values of the covariate. The regression slopes are heterogeneous, and each treatment group has a different slope and intercept. (F) A treatment × covariate interaction in which the order of the treatment groups does not differ for different values of the covariate. Both the treatment and the treatment × covariate interaction are significant.

and intercept, to each treatment group (Figure 10.4E,F). When the treatment × covariate interaction is significant, it may not be possible to discuss a general treatment effect because the difference among the treatments may depend on the value of the covariate. If the interaction is strong and the regression lines cross, the ordering of the treatment means will be reversed at high and low values of the covariate (Figure 10.4E). If the treatment effect is strong, the rank order of the treatment means may remain the same for different values of the covariate, even though the interaction term is significant (Figure 10.4F).

Comparing Means In the previous section, we emphasized that comparing the means of different treatment groups is essential for correctly interpreting the analysis of variance results. But how do we decide which means are truly different from one another? The ANOVA only tests the null hypothesis that the treatment means were all sampled from the same distribution. If we reject this null hypothesis, the ANOVA results do not specify which particular means differ from one another. To compare different means, there are two general approaches. With a posteriori (“after the fact”) comparisons, we use a test to compare all possible pairs of treatment means to determine which ones are different from one another. With a priori (“before the fact”) contrasts, we specify ahead of time particular combinations of means that we want to test. These combinations often reflect specific hypotheses that we are interested in assessing. Although a priori contrasts are not used often by ecologists and environmental scientists, we favor them for two reasons: first, because they are more

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specific, they are usually more powerful than generalized tests of pairwise differences of means. Second, the use of a priori contrasts forces an investigator to think clearly about which particular treatment differences are of interest, and how those relate to the hypotheses being addressed. We illustrate both a priori and a posteriori approaches with the analysis of a simple ANOVA design. The data come from Ellison et al. (1996), who studied the interaction between red mangrove (Rhizophora mangle) roots and epibenthic sponges. Mangroves are among the few vascular plants that can grow in fullstrength salt water; in protected tropical swamps, they form dense forests fringing the shore. The mangrove prop roots extend down to the substrate and are colonized by a number of species of sponges, barnacles, algae, and smaller invertebrates and microbes. The animal assemblage obviously benefits from this available hard substrate, but are there any effects on the plant? Ellison et al. (1996) wanted to determine experimentally whether there were positive effects of two common sponge species on the root growth of Rhizophora mangle. They established four treatments, with 14 to 21 replicates per treatment.9 The treatments were as follows: (1) unmanipulated; (2) foam attached to bare mangrove roots (the foam is a control “fake sponge” that mimics the hydrodynamic and other physical effects of a living sponge, but is biologically inert10); (3) Tedania ignis (red fire sponge) living colonies transplanted to bare mangrove roots; (4) Haliclona implexiformis (purple sponge) living colonies transplanted to bare mangrove roots. Replicates were established by randomly pre-selecting bare mangrove roots. Treatments were then randomly assigned to the replicates. The response variable was mangrove root growth, measured as mm/day for each 9

The complete design was actually a randomized block with unequal sample sizes, but we will treat it as a simple one-way ANOVA to illustrate the comparisons of means. For our simplified example, we analyze only 14 randomly chosen replicates for each treatment, so our analysis is completely balanced. 10

This particular foam control deserves special mention. In this book, we have discussed experimental controls and sham treatments that account for handling effects, and other experimental artifacts that we want to distinguish from meaningful biological effects. In this case, the foam control is used in a subtly different way. It controls for the physical (e.g., hydrodynamic) effects of a sponge body that disrupts current flow and potentially affects mangrove root growth. Therefore, the foam control does not really control for any handling artifacts because replicates in all four treatments were measured and handled in the same way. Instead, this treatment allows us to partition the effect of living sponge colonies (which secrete and take up a variety of minerals and organic compounds) from the effect of an attached sponge-shaped structure (which is biologically inert, but nevertheless alters hydrodynamics).

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Comparing Means

337

TABLE 10.12 Analysis of variance table for experiment testing effects of sponges on growth of red mangrove roots Degrees of freedom (df)

Source

Treatments Residual Total

3 52 55

Sum of Mean squares (SS) square (MS)

2.602 8.551 11.153

0.867 0.164

F-ratio

P-value

5.286

0.003

Four treatments were established (unmanipulated, foam, and two living sponge treatments) with 14 replicates in each treatment used in this analysis; total sample size = 14 × 4 = 56. The F-ratio for the treatment effect is highly significant, indicating that mean growth rates of mangrove roots differed significantly among the four treatments. Means and standard deviations of root growth (mm/day) for each treatment are illustrated in Figure 10.6. (Data from Ellison et al. 1996.)

replicate. Table 10.12 gives the analysis of variance for these data and Figure 10.5 illustrates the patterns among the means. The P-value for the F-ratio is highly significant (F3,52 = 5.286, P = 0.001), and the treatments account for 19% of the variation in the data (computed using Equation 10.23). The next step is to focus on which particular treatment means are different from one another. A Posteriori Comparisons

We will use Tukey’s “honestly significant difference” (HSD) test to compare the four treatment means in the mangrove growth experiment. This is one of many

Root growth rate (mm/day)

1.75 1.50 1.25 1.00 0.75 0.50 0.25 0.00 Unmanipulated Foam

Haliclona

Experimental treatment

Tedania

Figure 10.5 Growth rate (mm/day) of mangrove roots in four experimental treatments. Four treatments were established (unmanipulated, foam, and two living sponge treatments), with 14 replicates in each treatment. The height of the bar is the average growth rate for the treatment, and the vertical error bar is one standard deviation around the mean. The horizontal lines join treatment groups that do not differ significantly by Tukey’s HSD test (see Table 10.13). (Data from Ellison et al. 1996. Statistical analyses of these data are presented in Tables 10.12–10.15.)

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a posteriori procedures available for comparing pairs of means after ANOVA.11 Tukey’s HSD test statistically controls for the fact that we are carrying out many simultaneous comparisons. Therefore, the P-value must be adjusted downward for each individual test to achieve an experiment-wise error rate of α = 0.05. In the next section, we will discuss in more detail the general problem of how to deal with multiple P-values in a study. The first step is to calculate the HSD: ⎛1 1⎞ HSD = q ⎜ + ⎟ MSresidual ⎝ ni n j ⎠

(10.27)

where q is the value from a statistical table of the studentized range distribution, ni and nj are the sample sizes for the means of group i and group j that are being compared, and MSresidual is the familiar residual mean square from the one-way ANOVA table. For the data in Table 10.12, q = 3.42 (taken from a set of statistical tables), ni and nj = 14 replicates for all of the means, and MSresidual = 0.164. The calculated HSD = 0.523. Therefore, any pair of means among the four treatments that differs by at least 0.523 in average daily root growth is significantly different at P = 0.05. Table 10.13 shows the matrix of pair-wise differences between each of the means, and the corresponding P-value calculated by Tukey’s test. The analysis suggests that the unmanipulated roots differed significantly from growth of the roots with sponges attached. The two living sponge treatments did not differ significantly from each other or from the foam treatment. However, the foam treatment was marginally non-significant (P = 0.07) in comparison with the unmanipulated roots. These patterns are illustrated graphically in Figure 10.5, in which horizontal lines are placed over sets of treatment means that do not differ significantly from one another. Although these pair-wise tests did reveal particular pairs of treatments that differ, Tukey’s HSD test and other a posteriori tests may occasionally indicate that none of the pairs of means are significantly different from one another, even though the overall F-ratio led you to reject the null hypothesis! This inconsis11

Day and Quinn (1989) thoroughly discuss the different tests after ANOVA and their relative strengths and weaknesses. None of the alternatives is entirely satisfactory. Some suffer from excessive Type I or Type II errors, and others are sensitive to unequal sample sizes or variance differences among groups. Tukey’s HSD does control for multiple comparisons, although it may be slightly conservative with a potential risk of a Type II error (not rejecting H0 when it is false). See also Quinn and Keough (2002) for a discussion of the different choices.

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Comparing Means

TABLE 10.13 Mean differences between all possible pairs of treatments in an experimental study of effects of sponges on root growth of red mangrove

Unmanipulated Foam Haliclona Tedania

Unmanipulated

Foam

Haliclona

0.000 0.383 (0.072) 0.536 (0.031) 0.584 (0.002)

0.000 0.053 (0.986) 0.202 (0.557)

0.000 0.149 (0.766)

Tedania

0.000

Four treatments were established (unmanipulated, foam, two living sponge treatments) with 14 replicates in each treatment used in this analysis; total sample size = 14 × 4 = 56. The ANOVA is given in Table 10.12. This table illustrates the differences among the treatment means. Each entry is the difference in mean root growth (mm/day) of a pair of treatments. The diagonal of the matrix compares each treatment to itself, so the difference is always zero. The value in parentheses is the associated tail probability based on Tukey’s HSD test. This test controls the α level to account for the fact that six pairwise tests have been conducted. The larger the difference between the pairs of means, the lower the P-value. The unmanipulated roots differ significantly from the two treatments with living sponges (Tedania, Haliclona). Differences between all other pairs of means are non-significant. However, the pairwise comparison of the unmanipulated and foam treatments is only marginally above the P = 0.05 level, whereas the foam treatment does not differ significantly from either of the living sponge treatments. These patterns are illustrated graphically in Figure 10.5. (Data from Ellison et al. 1996.)

tent result can arise because the pairwise tests are not as powerful as the overall F-ratio itself. A Priori Contrasts

A priori contrasts are much more powerful, both statistically, and logically, than pairwise a posteriori tests. The idea is to establish contrasts, or specified comparisons between particular sets of means that test specific hypotheses. If we follow certain mathematical rules, a set of contrasts will be orthogonal or independent of one another, and they will actually represent a mathematical partitioning of the among-groups sum of squares. To create a contrast, we first assign an integer (positive, negative, or 0) to each treatment group. This set of integer coefficients is the contrast that we are testing. Here are the rules for building contrasts: 1. The sum of the coefficients for a particular contrast must equal 0. 2. Groups of means that are to be averaged together are assigned the same coefficient. 3. Means that are not included in the comparison of a particular contrast are assigned a coefficient of 0.

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Let’s try this for the mangrove experiment. If living sponge tissue enhances root growth, then the average growth of the two living sponge treatments should be greater than the growth of roots in the inert foam treatment. Our contrast values are Contrast I

control (0) foam (2) Tedania (–1)

Haliclona (–1)

Haliclona and Tedania receive the same coefficient (–1) because we want to compare the average of those two treatments with the foam treatment. The control receives a coefficient of 0 because we are testing an hypothesis about the effect of living sponge, so the relevant comparison is only with the foam. Foam receives a coefficient of 2 so that it is balanced against the average of the two living sponges and the sum of the coefficients is 0. Equivalently, you could have assigned coefficients of control (0), foam (6), Tedania (–3) and Haliclona (–3) to achieve the same contrast. Once the contrast is established, we use it to construct a new mean square, which has one degree of freedom associated with it. You can think of this as a weighted mean square, for which the weights are the coefficients that reflect the hypothesis. The mean square for each contrast is calculated as

MScontrast =

⎛ a ⎞ n ⎜ c iYi ⎟ ⎝ i =1 ⎠

∑

(10.28)

∑

c i2

i =1

The term in parentheses is just the sum of each coefficient (ci) multiplied by the – corresponding group mean (Yi). For our first contrast, the mean square for foam versus living sponge is MS foam vs living =

((0)(0.329) + (2)(0.712) + (−1)(0..765) + (− 1)(0.914)) 0 + 2 + ( − 1) + ( − 1) 2

× 14

= 0.151

(10.29)

This mean square has 1 degree of freedom, and we test it against the error mean square. The F-ratio is 0.151/0.164 = 0.921, with an associated P-value of 0.342 (Table 10.14). This contrast suggests that the growth rate of mangrove roots covered with foam was comparable to growth rates of mangrove roots covered with living sponges. This result makes sense because the means for these three treatment groups are fairly similar. There is no limit to the number of potential contrasts we could create. However, we would like our contrasts to be orthogonal to one another, so that the

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Comparing Means

results are logically independent. Orthogonal contrasts ensure that the P-values are not excessively inflated or correlated with one another. In order to create orthogonal contrasts, two additional rules must be followed: 4. If there are a treatment groups, at most there can be (a – 1) orthogonal contrasts created (although there are many possible sets of such orthogonal contrasts). 5. All of the pair-wise cross products must sum to zero (see Appendix). In other words, a pair of contrasts Q and R is independent if the sum of the products of their coefficients cQi and cRi equals zero: a

∑ cQic Ri = 0

(10.30)

i=1

Building orthogonal contrasts is a bit like solving a crossword puzzle. Once the first contrast is established, it constrains the possibilities for those that are remaining. For the mangrove root data, we can build two additional orthogonal contrasts to go with the first one. The second contrast is: Contrast II control (3) foam (–1) Tedania (–1)

Haliclona (–1)

This contrast sums to zero: it is orthogonal with the first contrast because it satisfies the cross products rule (Equation 10.30): (0)(3) + (2)(−1) + (−1)(−1) + (−1)(−1) = 0

(10.31)

This second contrast compares root growth in the control with the average of root growth of the foam and the two living sponges. This contrast tests whether enhanced root growth reflects properties of living sponges or the physical consequences of an attached structure per se. This contrast gives a very large F-ratio (14.00) that is highly significant (see Table 10.14). Again, this result makes sense because the average growth of the unmanipulated roots (0.329) is substantially lower than that of the foam treatment (0.712) or either of the two living sponge treatments (0.765 and 0.914). One final orthogonal contrast can be created: Contrast III control (0) foam (0)

Tedania (1)

Haliclona (–1)

The coefficients of this contrast again sum to zero, and the contrast is orthogonal to the first two because the sum of the cross products is zero (Equation 10.30). This third contrast tests for differences in growth rate of the two living sponge species but does not assess the mutualism hypothesis. It is of less inter-

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est than the other two contrasts, but it is the only contrast that can be created that is orthogonal to the first two. This contrast yields an F-ratio of only 0.947, with a corresponding P-value of 0.335 (see Table 10.14). Once again, this result is expected because the average growth rates of roots in the two living sponge treatments are similar (0.765 and 0.914). Notice that the sum of squares for each of these three contrasts add up to the total for the treatment sum of squares: SStreatment = SScontrast I + SScontrast II + SScontrast III 2 . 602

0 . 151 +

2 . 296

+ 0 . 155

(10.32)

In other words, we have now partitioned the total among-group sum of squares into three orthogonal, independent contrasts. This is similar to what happens in a two-way ANOVA, in which we decompose the overall treatment sum of squares into two main effects and an interaction term. This set of three contrasts is not the only possibility. We could have also specified this set of orthogonal contrasts: Contrast I control (1) foam (1) Tedania (–1) Contrast II control (1) foam (–1) Tedania (0) Contrast III control (0) foam (0) Tedania (1)

Haliclona (–1) Haliclona (0) Haliclona (–1)

Contrast I compares the average growth of roots covered with the living sponges with the average growth of roots in the two treatments with no living sponges. Contrast II specifically compares the growth of control roots versus foam-covered roots, and Contrast III compares growth rates of roots covered with Haliclona versus roots covered with Tedania. These results (Table 10.15) suggest that living sponges enhance root growth compared to unmanipulated roots or those with attached foam. However, the Contrast II reveals that root growth with foam was enhanced compared to unmanipulated controls, suggesting that mangrove root growth is responsive to hydrodynamics or some other physical property of attached sponges, rather than the presence of living sponge tissue. Either set of contrasts is acceptable, although we think the first set more directly addresses the effects of hydrodynamics versus living sponges. Finally, we note that there are at least two non-orthogonal contrasts that may be of interest. These would be control (0) control (0)

foam (1) foam (1)

Tedania (–1) Tedania (0)

Haliclona (0) Haliclona (–1)

These contrasts compare root growth with each of the living sponges versus root growth with the foam control. Both contrasts are non-significant (Tedania F1,52 = 0.12, P = 0.73; Haliclona F1,52 = 0.95, P = 0.33), again suggesting that the

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Comparing Means

effect of living sponges on mangrove root growth is virtually the same as the effect of biologically inert foam. However, these two contrasts are not orthogonal to one another (or to our other contrasts) because their cross products do not sum to zero. Therefore, these calculated P-values are not completely independent of the P-values we calculated in the other two sets of contrasts. In this example, the a posteriori comparisons using Tukey’s HSD test gave a somewhat ambiguous result, because it wasn’t possible to separate cleanly the mean values on a pairwise basis (see Table 10.13). The a priori contrasts (in contrast) gave a much cleaner result, with a clear rejection of the null hypothesis and confirmation that the sponge effect could be attributed to the physical effects of the sponge colony, and not necessarily to any biological effects of living sponge (see Tables 10.14 and 10.15). The a priori contrasts were more successful because they are more specific and more powerful than the pairwise tests. Notice also that the pairwise analysis required six tests, whereas the contrasts used only three specified comparisons. One cautionary note: a priori contrasts really do have to be established a priori—that is, before you have examined the patterns of the treatment means! If the contrasts are set up by grouping together treatments with similar means, the resulting P-values are entirely bogus, and the chance of a Type I error (falsely TABLE 10.14 A priori contrasts for analysis of variance for effects of sponges on mangrove root growth Source

Treatments

Foam vs. living Unmanipulated vs. living, foam Tedania vs. Haliclona Residual Total

Degrees of freedom (df)

Sum of Mean squares (SS) square (MS)

F-ratio

P-value

2.602

0.867

5.287

0.026

1 1

0.151 2.296

0.921 14.000

0.342 0.05, they are not considered significant by conventional hypothesis testing in the sciences. Thus, we would

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Tests for Goodness of Fit

conclude that our observed data fit a binomial distribution, and there is (just barely) insufficient evidence to reject the null hypothesis that the Belgian Euro is a fair coin.8 The classical, asymptotic tests and the exact test fail to reject only marginally the null hypothesis that the Euro coin is fair, testing the hypothesis P(data|H0). A Bayesian analysis instead tests P(H1| data) by asking: do the data actually provide evidence that the Belgian Euro is biased? To assess this evidence, we must calculate the posterior odds ratio, as described in Chapter 9 (using Equation 9.49). MacKay (2002) and Hamaker (2002) conducted these analyses using several different prior probability distributions. First, if we have no reason to initially prefer one hypothesis over another (H0: the coin is fair; H1: the coin is biased), then the ratio of their priors = 1 and the posterior odds ratio is equal to the likelihood ratio BAYESIAN ALTERNATIVE

P (data | H 0 ) P (data | H1) The likelihoods (numerator and denominator) of this ratio are calculated as: 1

P(D | H i ) = ∫ P(D | p, H i )P( p | H i )dp

(11.27)

where D is data, Hi is either hypothesis (i = 0 or 1), and p is the probability of success in a binomial trial. For the null hypothesis (fair coin), our prior probability is that there is no bias; thus p = 0.5, and the likelihood P(data |H0) is the probability of obtaining exactly 140 heads in 250 trials, = 0.0084 (see Footnote 8). For the alternative hypothesis (biased coin), however, there are at least three ways to set the prior.

This probability also can be calculated exactly using the binomial distribution. From Equation 2.3, the probability of getting exactly 140 heads in 250 spins with a fair coin is ⎛ 250⎞ 140 250−140 = 0.0084 ⎜ ⎟ 0.5 0.5 ⎝ 140⎠

But for a two-tailed significance test, we need the probability of obtaining 140 or more heads in our sample plus the probability of obtaining 110 or fewer tails in our sample. By analogy with the Fisher’s Exact Test (see Equation 11.13) we simply add up the probabilities of obtaining 140, 141, …, 250 tails and 0, 1, …, 110 tails in a sample of 250, with expected probability of 0.50. This sum = 0.0581, a value almost identical to that obtained with the chi-square and G-tests.

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First, we could use a uniform prior, which means that we have no initial knowledge as to how much bias there is and thus all biases are equally likely, and P(p|H1) = 1. The integral then reduces to the sum of the probabilities over all choices of p (from 0 to 1) of the binomial expansion for 140 heads out of 250 trials: 1 ⎛ 250 ⎞ 140 110 P( D | H 1 ) = ∑ ⎜ ⎟ p (1 − p) = 0.003 140 ⎝ ⎠ p=0

(11.28)

The likelihood ratio is therefore 0.0084/0.00398 = 2.1, or approximately 2:1 odds in favor of the Belgian Euro being a fair coin! Alternatively, we could use a more informative prior. The conjugate prior (see Footnote 11 in Chapter 5) for the binomial distribution is the beta distribution, P ( p | H1 , α ) =

Γ(2α) Γ(α)

p α −1(1 − p)α −1

(11.29)

where p is the probability of success, Γ(α) is the gamma distribution , and α is a variable parameter that expresses our prior belief in the bias of the coin. As α increases, our prior belief in the bias also increases. The likelihood of H1 is now P( D | H 1 ) =

Γ(2α) ⎛ 250 ⎞ Γ(α)2 ⎜⎝ 140 ⎟⎠

1 140+α−1

∫0 p

(1 − p)110+α−1dp

(11.30)

For α = 1 we have the uniform prior (and use Equation 11.28). By iteratively solving Equation 11.30, we can obtain likelihood ratios for a wide range of values of α. The most extreme odds-ratio obtainable using Equation 11.30 (remember, the numerator is fixed at 0.0084) is 0.52 (when α = 47.9). For this informative prior, the odds are approximately 2:1 (= 1/0.52) in favor of the Belgian Euro being a biased coin. Lastly, we could specify a very sharp prior that exactly matches the data: p = 140/250 = 0.56. Now, the likelihood of H1 = 0.05078 (the binomial expansion in Equation 11.28 with p = 0.56) and the likelihood ratio = 0.0084/0.05078 = 0.16 or 6:1 odds (= 1/0.16) in favor of the Belgian Euro having exactly the bias observed. Although our more informative priors suggest that the coin is biased, in order to reach that conclusion we had to specify priors that suggest strong advance knowledge that the Euro is indeed biased. An “objective” Bayesian analysis lets the data “speak for themselves” by using less informative priors, such as the uniform prior and Equation 11.28. That analysis provides little support for the hypothesis that the Belgian Euro is biased.

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Tests for Goodness of Fit

In sum, the asymptotic, exact, and Bayesian analyses all agree: the Belgian Euro is most likely a fair coin. The Bayesian analysis is very powerful in this context, because it provides a measure of the probability of the hypothesis of interest. In contrast, the conclusion of the frequentist analysis rests on P-values that are only slightly larger than the critical value of 0.05. GENERALIZING THE CHI-SQUARE AND G-TESTS FOR MULTINOMIAL DISCRETE DISTRIBU-

Both the chi-square and G-tests can be used to test the fit of data that have more than two levels to discrete distributions, such as the Poisson or multinomial. The procedure is the same: calculate expected values, apply Equation 11.5 or 11.7, and compute the tail probability of the test statistic relative to a χ2 distribution with ν degrees of freedom. The only tricky part is determining the degrees of freedom. We distinguish two types of distributions that we use in applying goodnessof-fit tests: distributions with parameters estimated independently of the data and distributions with parameters estimated from the data themselves. Sokal and Rohlf (1995) call these extrinsic hypotheses and intrinsic hypotheses, respectively. For an extrinsic hypothesis, the expectations are generated by a model or prediction that does not depend on the data themselves. Examples of extrinsic hypotheses include the 50:50 expectation for an unbiased Euro coin, and the 3:1 ratio of dominant to recessive phenotypes in a simple Mendelian cross of two heterozygotes. For extrinsic hypotheses, the degrees of freedom are always one less than the number of levels the variable can assume. Intrinsic hypotheses are those in which the parameters have to be estimated from the data themselves. An example of an intrinsic hypothesis is fitting data of counts of rare events to a Poisson distribution (see Chapter 2). In this case, the parameter of the Poisson distribution is not known, and must be estimated from the data themselves. We therefore use a degree of freedom for the total number of individuals sampled and use another degree of freedom for each estimated parameter. Because the Poisson distribution has one additional parameter (λ, the rate parameter), the degrees of freedom for that test would be two less than the number of levels. It also is possible to use a chi-square or G-test to test the fit of a dataset to a normal distribution, but in order to do that, the data first have to be grouped into discrete levels. The normal distribution has two parameters to be estimated from the data (the mean μ and the standard deviation σ). Therefore, the degrees of freedom for a chi-square or G-test for goodness-of-fit to a normal distribution equals three less than the number of levels. However, the normal distribution is continuous, not discrete, and we would have to divide the data up into an arbitrary number of discrete levels in order to conduct the test. The TIONS

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results of such a test will differ depending on the number of levels chosen. To assess the goodness-of-fit for continuous distributions, the Kolmogorov-Smirnov test is a more appropriate and powerful test. Testing Goodness-of-Fit for Continuous Distributions: The Kolmogorov-Smirnov Test

Many statistical tests require that either the data or the residuals fit a normal distribution. In Chapter 9, for example, we illustrated diagnostic plots for examining the residuals from a linear regression (see Figures 9.5 and 9.6). Although we asserted that Figure 9.5A illustrated the expected distribution of residuals for a linear model with a normal distribution of errors, we would like to have a quantitative test for that assertion. The Kolmogorov-Smirnov test is one such test. The Kolmogorov-Smirnov test compares the distribution of an empirical sample of data with an hypothesized distribution, such as the normal distribution. Specifically, this test compares the empirical and expected cumulative distribution functions (CDF). The CDF is defined as the function F(Y) = P(X < Y) for a random variable X. In words, if X is a random variable with a probability density function (PDF) f(X), then the cumulative distribution function F(Y) equals the area under f(X) in the interval X < Y (see Chapter 2 for further discussion of PDF’s and CDF’s). Throughout this book we have used the cumulative distribution function extensively—the tail probability, or P-value, is the area under the curve beyond the point Y, which is equal to 1 – F(Y). Figure 11.4 illustrates two CDF’s. The first is the empirical CDF of the residual values from the linear regression described in Chapter 9 of log10(plant species richness) on log10(island area). The second is the hypothetical CDF that would be expected if these residuals were normally distributed with mean = 0 and standard deviation = 0.31 (these values are estimated from the residuals themselves). The CDF of the empirical data is not a smooth curve because we have only 17 points, whereas the expected CDF is a smooth curve because it comes from an underlying normal distribution with infinitely many points. The Kolmogorov-Smirnov test is a two-sided test. The null hypothesis is that the observed CDF [Fobs(Y)] = the CDF of the hypothesized distribution [Fdist(Y)]. The test statistic is the absolute value of the maximum vertical difference between the observed and expected CDF’s (arrow in Figure 11.4). Basically, one measures this distance at each observed point and takes the largest one. For Figure 11.4, the maximum distance is 0.148. This maximum difference is compared with a table of critical values (e.g., Lilliefors 1967) for a particular sample size and P-value. If the maximum difference is less than the associated critical value, we cannot reject the null hypothesis that our data do not differ

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Tests for Goodness of Fit

1.0

Cumulative probability

0.8

0.6

0.4

0.2

Test statistic = maximum difference between observed and expected

0.0 –0.5

0.0

0.5 Residual

Figure 11.4 Goodness-of-fit test for a continuous distribution. Comparison of the cumulative distribution function of the observed residuals (black points, gray line) and the expected distribution if they were from a normal distribution (blue line). Residuals are calculated from the species-area regression of log10(species number) on log10(island area) for plant species of the Galápagos (data in Table 8.2; see Figure 9.6). The KolmogorovSmirnov test for goodness-of-fit compares these two distributions. The test statistic is the maximum difference between these two distributions, indicated by the double-headed arrow. The maximum difference for these data is 0.148, compared to a critical value of 0.206 for a test at P = 0.05. Because the null hypothesis cannot be rejected, the residuals appear to follow a normal distribution, which is one of the assumptions of linear regression (see Chapter 9). The Kolmogorov-Smirnov test can be used to compare a sample of continuous data to any continuous distribution, such as the normal, exponential, or log-normal (see Chapter 2).

from the expected distribution. In our example, the critical value for P = 0.05 is 0.206. Because the observed maximum difference was only 0.148, we conclude that the distribution of residuals from our regression is consistent with the hypothesis that the true errors come from a normal distribution (we do not reject H0). On the other hand, if the maximum difference had been greater than the 0.206, we would reject the null hypothesis, and we should question whether a linear regression model, which assumes that the errors have a normal distribution, is the appropriate one to use for analyzing the data. The Kolmogorov-Smirnov test is not limited to the normal distribution; it can be used for any continuous distribution. If you want to test whether your data are log-normally or exponentially distributed, for example, you can use the Kolmogorov-Smirnov test to compare the empirical CDF with a log-normal or exponential CDF (Lilliefors 1969).

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Summary Categorical data from tabular designs are a common outcome of ecological and environmental research. Such data can be analyzed easily using the familiar chisquare or G-tests. Hierarchical log-linear models or classification trees can test detailed hypotheses regarding associations among and between categorical variables. A subset of these methods also can be used to test whether or not the distribution of the data and residuals are consistent with, or are fit by, particular theoretical distributions. Bayesian alternatives to these tests have been developed, and these methods allow for estimation of expected frequency distributions and parameters of the log-linear models.

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CHAPTER 12

The Analysis of Multivariate Data

All of the analytical methods that we have described so far apply only to single response variables, or univariate data. Many ecological and environmental studies, however, generate two or more response variables. In particular, we often analyze how multiple response variables—multivariate data—are related simultaneously to one or more predictor variables. For example, a univariate analysis of pitcher plant size might be based on a single variable: pitcher height. A multivariate analysis would be based on multiple variables: pitcher height, mouth opening, tube and keel widths, and wing lengths and spread (see Table 12.1). Because these response variables are all measured on the same individual, they are not independent of one another. Statistical methods for analyzing univariate data—regression, ANOVA, chi-square tests, and the like—may be inappropriate for analyzing multivariate data. In this chapter, we introduce several classes of methods that ecologists and environmental scientists use to describe and analyze multivariate data. As with the univariate methods we discussed in Chapters 9–11, we can only scratch the surface and describe the important elements of the most common forms of multivariate analysis. Gauch (1982) and Manly (1991) present classical multivariate analysis techniques used commonly by ecologists and environmental scientists, whereas Legendre and Legendre (1998) provide a more thorough treatment of developments in multivariate analysis through the mid-1990s. Development of new multivariate methods as well as assessment of existing techniques is an active area of statistical research. Throughout this chapter, we highlight some of these newer methods and the debates surrounding them.

Approaching Multivariate Data Multivariate data look very much like univariate data: they consist of one or more independent (predictor) variables and two or more dependent (response)

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variables. The distinction between univariate and multivariate data largely lies in how the data are organized and analyzed, not in how they are collected.1 Most ecological and environmental studies yield multivariate data. Examples include a set of different morphological, allometric, and physiological measurements taken on each of several plants assigned to different treatment groups; a list of species and their abundances recorded at multiple sampling stations along a river; a set of environmental variables (e.g., temperature, humidity, rainfall, etc.) and a corresponding set of organismal abundances or traits measured at several sites across a geographic gradient. In some cases, however, the explicit goal of a multivariate analysis can alter the sampling design. For example, a mark–recapture study of multiple species at multiple sites would not be designed the same way as a mark–recapture study of a single species because different methods of analysis might have to be used for rare versus common species (see Chapter 14). Multivariate data can be quantitative or qualitative, continuous, ordered, or categorical. We focus our attention in this chapter on continuous quantitative variables. Extensions to qualitative multivariate variables are discussed by Gifi (1990). The Need for Matrix Algebra

The mathematics of multivariate analysis is expressed best using matrix algebra. Matrix algebra allows us to use equations for multivariate analysis that look like those we have used earlier for univariate analysis. The similarity is more than skin deep; the interpretations of these equations are close, and many of the equations used for univariate statistical methods also can be written using matrix notation. We summarize basic matrix notation and matrix algebra in the Appendix. A multivariate random variable Y is a set of n univariate variables {Y1, Y2, Y3, …, Yn} that are all measured for the same observational or experimental unit such as a single island, a single plant, or a single sampling station. We designate a multivariate random variable as a boldfaced, capital letter, whereas we con-

In fact, you have already been exposed to multivariate data in Chapters 6, 7, and 10. In a repeated-measures design, multiple observations are taken on the same individual at different times. In a randomized block or split-plot design, certain treatments are physically grouped together in the same block. Because the response variables measured within a block or on a single individual are not independent of one another, the analysis of variance has to be modified to take account of this data structure (see Chapter 10). In this chapter, we will describe some multivariate methods in which there is not a corresponding univariate analysis.

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Approaching Multivariate Data

tinue to write univariate random variables as italicized, capital letters. Each observation of Y is written as yi , which is a row vector. The elements yi,j of this row vector correspond to the measurement of the jth variable Yj ( j ranges from 1 to n variables) measured for individual i (i ranges from 1 to m individual observations or experimental units): yi = [yi,1, yi,2 , yi,3 , … yi,j , …, yi,n]

(12.1)

For convenience, we usually omit the subscript i in Equation 12.1. The row vector y is an example of a matrix (plural matrices) with one row and n columns. Each observation yi,j within the square brackets is called an element of the matrix (or row vector) y. An ecologist would organize typical multivariate data in a simple spreadsheet with m rows, each representing a different individual, and n columns, each representing a different measured variable. Each row of the matrix corresponds to the row vector yi, and the entire table is the matrix Y. An example of a multivariate dataset is presented in Table 12.1. Each of the m = 89 rows represents a different plant, and each of the n = 10 columns represents a different morphological variable that was measured for each plant. Several matrices will recur throughout this chapter. Equation 12.1 is the basic n-dimensional vector describing a single observation: y = [y1, y2, y3, …, yn]. We also define a vector of means as Y = [Y1 ,Y2 ,Y3 , …,Yn ]

(12.2)

TABLE 12.1 Multivariate data for a carnivorous plant, Darlingtonia californica Site

TJH TJH ...

Plant Height Mouth Tube Keel Wing1 Wing2 Wspread Hoodmass Tubemass Wingmass

1 2 ...

654 413 ...

38 22 ...

17 17 ...

6 6 ...

85 55 ...

76 26 ...

55 60 ...

1.38 0.49 ...

3.54 1.48 ...

0.29 0.06 ...

Each row represents a single observation of a cobra lily (Darlingtonia californica) plant measured at a particular site (T. J. Howell’s fen, TJH) in the Siskiyou Mountains of southern Oregon. Each column represents a different morphological variable. Measurements for each plant included seven morphological variables (all in mm): height (height), mouth opening (mouth), tube diameter (tube), keel diameter (keel), length of each of the two “wings” (wing1, wing2) making up the fishtail appendage, and distance between the two wing tips (wspread). The plants were dried and the hood (hoodmass), tube (tubemass), and fishtail appendage (wingmass) were weighed (± 0.01 g). The total dataset, collected in 2000 by A. Ellison, R. Emerson, and H. Steinhoff, consists of 89 plants measured at 4 sites. This is a typical multivariate dataset: the different variables measured on the same individual are not independent of one another.

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which is the n-dimensional vector of sample means (calculated as described in Chapter 3) of each Yj . The elements of this row vector are the means of each of – the n variables, Yj . In other words, Y is equal to the set of sample means of each column of our original data matrix Y. In a univariate analysis, the corresponding measurement is the sample mean of the single response variable, which is – an estimate of the true population mean μ. Similarly, the vector Y is an estimate of the vector m = [μ1, μ2, μ3, …, μn] of the true population means of each of the n variables. We need three matrices to describe the variation among the yi,j observations. The first is the variance-covariance matrix, C (see Footnote 5 in Chapter 9). The variance-covariance matrix is a square matrix, in which the diagonal elements are the sample variances of each variable (calculated in the usual way; see Chapter 3, Equation 3.9). The off-diagonal elements are the sample covariances between all possible pairs of variables (calculated as we did for regression and ANOVA; see Chapter 9, Equation 9.10). ⎡ s12 c12 ⎢ c s22 C = ⎢⎢ 2,1 ⎢ ⎢⎣c n,1 c n,2

… c1,n ⎤ ⎥ … c 2,n ⎥ ⎥ ⎥ … sn2 ⎥⎦

(12.3)

In Equation 12.3, sj2 is the sample variance of variable Yj , and cj,k is the sample covariance between variables Yj and Yk . Note that the sample covariance between variables Yj and Yk is the same as the sample covariance between variables Yk and Yj . Therefore, the elements of the variance-covariance matrix are symmetric, and are mirror images of one another across the diagonal (cj,k = ck,j). The second matrix we use to describe the variances is the matrix of sample standard deviations: ⎡ ⎢ ⎢ D(s) = ⎢ ⎢ ⎢ ⎣

s12

…

0 0

s22

… …

0 ⎤ ⎥ 0 ⎥ ⎥ ⎥ sn2 ⎥⎦

(12.4)

D(s) is a diagonal matrix: a matrix in which elements are all 0 except for those on the main diagonal. The non-zero elements are the sample standard deviations of each Yj.

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Comparing Multivariate Means

Lastly, we need a matrix of sample correlations: ⎡ 1 r12 ⎢ r2,1 1 P= ⎢ ⎢ ⎢ ⎢⎣rn,1 rn,2

… r1,n ⎤ ⎥ … r2,n ⎥ ⎥ ⎥ … 1 ⎥⎦

(12.5)

In this matrix, each element rj,k is the sample correlation (see Chapter 9, Equation 9.19) between variables Yj and Yk. Because the correlation of a variable with itself = 1, the diagonal elements of P all equal 1. Like the variance-covariance matrix, the matrix of correlations is symmetric (rj,k = rk,j).

Comparing Multivariate Means Comparing Multivariate Means of Two Samples: Hotelling’s T 2 Test

We introduce the concepts of multivariate analysis through a straightforward extension of the univariate t-test to the multivariate case. The classical t-test is used to test the null hypothesis that the mean value of a single variable does not differ between two groups or populations. For example, we could test the simple hypothesis that pitcher height of Darlingtonia californica (see Table 12.1) does not differ between populations sampled at Day’s Gulch (DG) and T.J. Howell’s fen (TJH). At each site, 25 plants were randomly sampled, from which we first calculated the mean height (DG = 618.8 mm; TJH = 610.4 mm) and the standard deviation (DG = 100.6 mm; TJH = 83.7 mm). Based on a standard ttest, the plants from these two populations do not differ significantly in height (t = 0.34, with 48 degrees of freedom, P = 0.74). We could conduct additional t-tests for each of the variables in Table 12.1 with or without corrections for multiple tests (see Chapter 10). However, we are more interested in asking whether overall plant morphology—quantified as the vector of means of all the morphological variables taken together—differs between the two sites. In other words, we wish to test the null hypothesis that the vectors of means of the two groups are equal. Hotelling’s T 2 test (Hotelling 1931) is a generalization of the univariate t-test to multivariate data. We first compute for each group the means of each of the variables Yj of interest and then assemble them into two mean column vectors – – (see Equation 12.2), Y1 (for TJH) and Y2 (for DG). Note that a column vector is simply a row vector that has been transposed—the columns and rows are interchanged. The rules of matrix algebra, explained in the Appendix, some-

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times require vectors to be single columns, and sometimes require vectors to be single rows. But the data are the same regardless of whether they are represented as row vectors or column vectors. For this example, we have seven morphological variables: pitcher height, mouth opening, tube and keel widths, and lengths and spread of the wings of the fishtail appendage at the mouth of the pitcher (see Table 12.1), each measured at two sites. ⎡610.0⎤ ⎥ ⎢ ⎢ 31.2 ⎥ ⎢ 19.9 ⎥ ⎥ ⎢ Y1 = ⎢ 6.7 ⎥ ⎢ 61.5 ⎥ ⎥ ⎢ ⎢ 59.9 ⎥ ⎢ 77.9 ⎥ ⎦ ⎣

⎡618.8⎤ ⎥ ⎢ ⎢ 33.1 ⎥ ⎢ 17.9 ⎥ ⎥ ⎢ Y2 = ⎢ 5.6 ⎥ ⎢ 82.4 ⎥ ⎥ ⎢ ⎢ 79.4 ⎥ ⎢ 84.2 ⎥ ⎦ ⎣

(12.6)

– For example, average pitcher height at TJH was 610.0 mm (the first element of Y1), whereas average pitcher mouth opening at DG was 33.1 mm (the second element – of Y2). Incidentally, the variables used in a multivariate analysis do not necessarily have to be measured in the same units, although many multivariate analyses rescale variables into standardized units, as described later in this chapter. Now we need some measure of variance. For the t-test, we used the sample variances (s2) from each group. For Hotelling’s T 2 test, we use the sample variance-covariance matrix C (see Equation 12.3) for each group. The sample variance-covariance matrix for site TJH is ⎡7011.5 284.5 32.3 −12.5 137.4 ⎢ ⎢ 284.5 31.8 −0.7 −1.9 55.8 ⎢ 32.3 −0.7 5.9 0.6 −19.9 ⎢ C1 = ⎢ −12.5 −1.9 0.6 1.1 −0.9 ⎢ 137.4 55.8 −19.9 −0.9 356.7 ⎢ −6.9 −3.0 305.6 ⎢ 691.7 77.7 ⎢ 76.5 66.3 −13.9 −5.6 397.4 ⎣

691.7 77.7 −6.9 −3.0 305.6 482.1 511.6

76.5 ⎤ ⎥ 66.3 ⎥ −13.9 ⎥ ⎥ −5.6 ⎥ 397.4 ⎥ ⎥ 511.6 ⎥ 973.3 ⎥⎦

(12.7)

This matrix summarizes all of the sample variances and covariances of the variables. For example, at TJH, the sample variance of pitcher height was 7011.5 mm2, whereas the sample covariance between pitcher height and pitcher mouth

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Comparing Multivariate Means

opening was 284.5 mm2. Although a sample variance-covariance matrix can be constructed for each group (C1 and C2), Hotelling’s T 2 test assumes that these two matrices are approximately equal, and uses a matrix CP that is the pooled estimate of the covariance of the two groups: CP =

[(m1 − 1)C1 + (m2 − 1)C 2] (m1 + m2 − 2)

(12.8)

where m1 and m2 are the sample sizes for each group (here, m1 = m2 = 25). Hotelling’s T 2 statistic is calculated as m1m2 (Y1 − Y2 ) C P −1(Y1 − Y2 ) (m1 + m2 ) T

T2=

(12.9)

– – where Y1 and Y2 are the vectors of means (see Equation 12.2), CP is the pooled sample variance-covariance matrix (see Equation 12.8), superscript T denotes matrix transpose, and superscript (–1) denotes matrix inversion (see Appendix). Note that in this equation, there are two types of multiplication. The first is matrix multiplication of the vectors of differences of means and the inverse of the covariance matrix: −1

(Y1 − Y2 )T C P (Y1 − Y2 )

The second is scalar multiplication of this product by the product of the sample sizes, m1m2. Following the rules of matrix multiplication, Equation 12.9 gives T 2 as a scalar, or a single number. A linear transformation of T 2 yields the familiar test statistic F: F=

(m1 + m2 − n − 1)T 2 (m1 + m2 − 2)n

(12.10)

where n is the number of variables. Under the null hypothesis that the population mean vectors of each group are equal (i.e., m1 = m2), F is distributed as an F random variable with n numerator and (m1 + m2 – n –1) denominator degrees of freedom. Hypothesis testing then proceeds in the usual way (see Chapter 5). For the seven morphological variables, T 2 equals 84.62, with 7 numerator and 42 denominator degrees of freedom. Applying Equation 12.10 gives an F-value of 10.58. The critical value of F with 7 numerator and 42 denominator degrees of freedom equals 2.23, which is much less than the observed value of 10.58.

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Therefore, we can reject the null hypothesis (with a P-value of 1.3 × 10–7) that the two population mean vectors, m1 and m2 , are equal. Comparing Multivariate Means of More Than Two Samples: A Simple MANOVA

If we wish to compare univariate means of more than two samples of treatment groups, we use ANOVA in lieu of the standard t-test. Similarly, if we are comparing multivariate means of more than two groups, we use a multivariate ANOVA, or MANOVA, in lieu of Hotelling’s T 2 test. As in Hotelling’s T 2 test, we have j = 1, …, n variables or measurements taken for each individual and i = 1, …, m total observations. In the MANOVA, however, we have k = 1, …, g groups, and l = 1, …, q observations within each group. The multivariate observations (Equation 12.1) are denoted as yk,l , the lth vector of observations within the kth group. If our design is balanced, so that there are the same number q of observations in each of the g groups, then our total sample size m = gq. By analogy with a one-way ANOVA (see Chapter 10, Equation 10.6), we analyze the model Y = m + Ak + ekl

(12.11)

where Y is the matrix of measurements (with m rows of observations and n columns of variables), m is the population (grand) mean, Ak is the matrix of deviations of the kth treatment from the sample grand mean, and ekl is the error term: the difference between the lth individual in the kth treatment group and the mean of the kth treatment. The ekl’s are assumed to come from a multivariate normal distribution (see next section). The procedure is basically the same as a one-way ANOVA (Chapter 10), except that instead of comparing the group means, we compare the group centroids—the multivariate means.2 The null hypothesis is that the means of the treatment groups are all the same: m1 = m2 = m3 = … = mg. As with ANOVA, the test statistic is the ratio of the among-group

In a one-dimensional space, two means can be compared as the arithmetic difference between them, which is what an ANOVA does for univariate data. In a two-dimensional space the mean vector of each group can be plotted as a point (x–, –y ) in a Cartesian graph. These points represent the centroids (often thought of as the center of gravity of the cloud of points), and we can calculate the geometric distance between them. Finally, in a space with three (or more) dimensions, the centroid can again be located by a set of Cartesian coordinates with one coordinate for each dimension in the space. MANOVA compares the distances among those centroids, and tests the null hypothesis that the distances among the group centroids are no more different from those expected due to random sampling.

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Comparing Multivariate Means

sums of squares divided by the within-group sums of squares, but these are calculated differently for MANOVA, as described below. This ratio has an F-distribution (see Chapter 10). In an ANOVA, the sums of squares are simple numbers; in a MANOVA, the sums of squares are matrices—called the sums of squares and cross-products (SSCP) matrices. These are square matrices (equal number of rows and columns) for which the diagonal elements are the sums of squares for each variable and for which the off-diagonal elements are the sums of cross-products for each pair of variables. SSCP matrices are directly analogous to sample variance-covariance matrices (see Equation 12.3), but we need three of them. The among-groups SSCP matrix is the matrix H: THE SSCP MATRICES

H = q ∑ (Yk. − Y.. )(Yk. − Y.. )T

(12.12)

k =1

— In Equation 12.12, Y k• is the vector of sample means in group k of the q observations in that group: q 1 Yk. = ∑ y kl q l =1 — and Y .. is the vector of sample means over all treatment groups g

Y.. =

1 kq k =1

∑ ∑ ykl l =1

The within-group SSCP matrix is the matrix E: g

E = ∑ ∑ (ykl − Yk. )(ykl − Yk. )T

(12.13)

k =1 l =1

— with Y k• again being the vector of sample means of treatment group k. In a univariate analysis, the analog of H in Equation 12.12 is the among-group sum of squares (see Equation 10.2, Chapter 10), and the analog of E in Equation 12.13 is the within-group sum of squares (Equation 10.3, Chapter 10). Lastly, we compute the total SSCP matrix: g

T = ∑ ∑ (y kl − Y.. )(y kl − Y.. )T k =1 l =1

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(12.14)

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Four test-statistics are generated from the H and E matrices: Wilk’s lambda, Pillai’s trace, Hotelling-Lawley’s trace, and Roy’s greatest root (see Scheiner 2001). These are calculated as follows: TEST STATISTICS

Wilk’s lambda = Λ =

E E+H

where | | denotes the determinant of a matrix (see Appendix).

∑ ⎜⎝ λ i +i 1⎟⎠ = trace [(E + H)−1 H] s

Pillai’s trace =

⎛ λ

⎞

i=1

where s is the smaller of either the degrees of freedom (number of groups g –1) or the number of variables n; λi is the ith eigenvalue (see Appendix Equation A.17) of E –1H; and trace is the trace of a matrix (see Appendix).

∑ λ i = trace (E −1 H) n

Hotelling-Lawley’s trace =

i=1

and Roy’s greatest root = θ =

λ1 λ1 + 1

where λ1 is the largest (first) eigenvalue of E –1H. For large sample sizes, Wilk’s lambda, Hotelling-Lawley’s trace, and Pillai’s trace all converge to the same P-value, although Pillai’s trace is the most forgiving of violations of assumptions such as multivariate normality. Most software packages will give all of these test statistics. Each of these four test statistics can be transformed into an F-ratio and tested as in ANOVA. However, their degrees of freedom are not the same. Wilk’s lambda, Pillai’s trace, and Hotelling-Lawley’s trace are computed from all of the eigenvalues and hence have a sample size of nm (number of variables n × number of samples m) with n(g – 1) numerator degrees of freedom (g is the number of groups). Roy’s largest root uses only the first eigenvalue, so its numerator degrees of freedom are only n. The degrees of freedom for Wilk’s lambda often are fractional, and the associated F-statistic is only an approximation (Harris 1985). The choice among these MANOVA test statistics is not critical. They usually give very similar results, as in the analysis of the Darlingtonia data (Table 12.2). As with ANOVA, if the MANOVA yields significant results, you may be interested in determining which particular groups dif-

COMPARISONS AMONG GROUPS

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TABLE 12.2 Results of a one-way MANOVA I. H matrix

Height Mouth Tube Keel Wing1 Wing2 Wspread

Height 35118.38 5584.16 –1219.21 –2111.58 17739.19 11322.59 18502.78

Mouth 5584.16 1161.14 –406.04 –395.59 2756.84 1483.22 1172.86

Tube –1219.21 –406.04 229.25 127.10 –842.68 –432.92 659.62

Keel –2111.58 –395.59 127.10 142.49 –1144.14 –706.07 –748.92

Wing1 17739.19 2756.84 –842.68 –1144.14 12426.23 9365.82 10659.12

Wing2 11322.59 1483.22 –432.92 –706.07 9365.82 7716.96 8950.57

Wspread 18502.78 1172.86 659.62 –748.92 10659.12 8950.57 21440.44

Height 834836.33 27526.88 8071.43 1617.17 37125.87 46657.82 18360.69

Mouth 27526.88 2200.17 324.05 –21.91 4255.27 4102.43 3635.34

Tube 8071.43 324.05 671.82 196.12 305.16 486.06 375.09

Keel 1617.17 –21.91 196.12 265.49 –219.06 –417.36 –632.27

Wing1 37125.87 4255.27 305.16 –219.06 31605.96 28737.10 33487.39

Wing2 46657.82 4102.43 486.06 –417.36 28737.10 39064.26 41713.77

Wspread 18360.69 3635.34 375.09 –632.27 33487.39 41713.77 86181.61

II. E matrix

Height Mouth Tube Keel Wing1 Wing2 Wspread

III. Test statistics

Statistic Pillai’s trace Wilk’s lambda Hotelling-Lawley’s trace Roy’s greatest root

Value 1.11 0.23 2.09 1.33

F 6.45 6.95 7.34 14.65

Numerator df 21 21 21 7

Denominator df 231 215.91 221 77

P-value 3 × 10–14 3 × 10–15 3 × 10–16 5 × 10–12

The original data are measurements of 7 morphological variables on 89 individuals of the cobra lily Darlingtonia californica collected at 4 sites (see Table 12.1). The H matrix is the among-groups sums of squares and crossproducts (SSCP) matrix. The diagonal elements are the sums of squares for each variable, and the off-diagonal elements are the sums of cross-products for each pair of variables (see Equation 12.12). The H matrix is analogous to the univariate calculation of the among-groups sum of squares (see Equation 10.2). The E matrix is the within-groups SSCP matrix. The diagonal elements are the within-group deviations for each variable, and the off-diagonal elements are the residual cross-products for each pair of variables (see Equation 12.13). The E matrix is analogous to the univariate calculation of the residual sum of squares (see Equation 10.3). In these matrices, we depart from the good practice of reporting only significant digits in order to avoid round-off error in our calculations. If we were to report these data in a scientific publication, they would not have more significant digits that we had in our measurements (see Table 12.1). The four test statistics (Pillai’s trace, Wilk’s lambda, Hotelling-Lawley’s trace, and Roy’s greatest root) represent different ways to test for differences among groups in a multivariate analysis of variance (MANOVA). All of these measures generated very small P-values, suggesting that the 7-element vector of morphological measurements for Darlingtonia differed significantly among sites.

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fer from one another. For post-hoc comparisons, Hotelling’s T2 test with the critical value adjusted using the Bonferroni correction (see Chapter 10) can be used for each pair-wise comparison. Discriminant analysis (described later in this chapter) also can be used to determine how well the groups can be separated. All of the ANOVA designs described in Chapter 10 have direct analogues in MANOVA when the response variables are multivariate. The only difference is that the SSCP matrices are used in place of the among- and within-group sums of squares. Computing SSCP matrices can be difficult for complex experimental designs (Harris 1985; Hand and Taylor 1987). Scheiner (2001) summarizes MANOVA techniques for ecologists and environmental scientists. ASSUMPTIONS OF MANOVA In addition to the usual assumptions of ANOVA (observations are independent and randomly sampled, and within-group errors are equal among groups and normally distributed), MANOVA has two additional assumptions. First, similar to the requirements of Hotelling’s T 2 test, the covariances are equal among groups (the assumption of sphericity). Second, the multivariate variables used in the analysis and the error terms ekl in Equation 12.11 must conform to a multivariate normal distribution. In the next section, we describe the multivariate normal distribution and how to test for departures from it. Because Pillai’s trace is robust to modest violations of these assumptions, in practice MANOVA is valid if the data do not depart substantially from sphericity or multivariate normality. Analysis of Similarity (ANOSIM) is an alternative method to MANOVA (Clarke and Green 1988; Clarke 1993) that does not depend on multivariate normality, but ANOSIM can be used only for one-way and fully crossed or nested two-way designs. ANOSIM has lower power (high probability of Type II statistical error) if strong gradients are present in the data (Somerfield et al. 2002).

The Multivariate Normal Distribution Most multivariate methods for testing hypotheses require that the data being analyzed conform to the multivariate normal (or multinormal) distribution. The multivariate normal distribution is the analog of the normal (or Gaussian) distribution for the multidimensional variable Y = [Y1, Y2, Y3, … , Yn]. As we saw in Chapter 2, the normal distribution has two parameters: μ (the mean) and σ2 (the variance). In contrast, the multivariate normal distribution3 is defined by the mean vector m and the covariance matrix S; the number of parameters in the multivariate normal distribution depends on the number of random variables Yi in Y. As you know, a one-dimensional normal distribution has a bell-

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The Multivariate Normal Distribution

shaped curve (see Figure 2.6). For a two-dimensional variable Y = [Y1, Y2], the bivariate normal distribution looks like a bell or hat (Figure 12.1). Most of the probability mass is concentrated near the center of the hat, closest to the means of the two variables. As we move in any direction toward the rim of the hat, the probability density becomes thinner and thinner. Although it is difficult to draw a multivariate distribution for variables of more than two dimensions, we can calculate and interpret them the same way. For most purposes, it is convenient to use the standardized multivariate normal distribution, which has a mean vector m = [0]. The bivariate normal distribution illustrated in Figure 12.1 is actually a standardized bivariate normal distribution. We also can illustrate this distribution as a contour plot (Figure 12.1E) of concentric ellipses. The central ellipse corresponds to a slice of the peak of the distribution, and as we move down the hat, the elliptical slices get larger. When all the variables Yn in Y are independent and uncorrelated, all the off-diagonal correlation coefficients rmn, in the correlation matrix P = 0, and the elliptical slices are perfectly circular. However, variables in most multivariate data sets are correlated, so the slices usually are elliptical in shape, as they are in Figure 12.1E. The stronger the correlation between a pair of variables, the more drawn out are the contours of the ellipse. The idea can be extended to more than two variables in Y, in which case the slices would be ellipsoids or hyperellipsoids in multi-dimensional space. 3 The probability density function for a multivariate normal distribution assumes an n-dimensional random vector Y of random variables [Y1, Y2, …, Yn] with mean vector m = [μ1, μ2, …, μn] and variance-covariance matrix

γ 1,n ⎤ ⎥ γ 2,n ⎥ ⎥ ⎥ ⎥ σ n2 ⎥ ⎦

⎡σ2 γ 12 ⎢ 1 ⎢γ σ2 ∑ = ⎢ 2,1 2 ⎢ ⎢ ⎢⎣ γ n,1 γ n,2

in which σi2 is the variance of Yi and γij is the covariance of Yi and Yj. For this vector Y, the probability density function for the multivariate normal distribution is f (Y ) =

1 (π )n ∑

⎡ 1 T − 1(Y −μ ) ⎤ ⎢ − 2 ( Y −μ ) ∑ ⎥ ⎦

e⎣

See the Appendix for details on matrix determinants (| |), inverses (matrices raised to the (–1) power) and transposition (T). The number of parameters needed to define the multivariate normal distribution is 2n + n(n – 1)/2, where n is the number of variables in Y. Notice how this equation looks like the probability density function for the univariate normal distribution (see Footnote 10, Chapter 2, and Table 2.4).

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(A)

(B)

0.4

0.2 0.1

1 X2

Probability

0.3 0.2

0.0

–2

0 0 X1

–3 –3 –2 –1

0 X2

–3 –2 –1

(D)

0 X1

(E) 0.25

0.15

0.05

1 X2

–3 –2 –1

0 –1

0.1

Probability

0.3

2 2

X 0 2

–2

0 –2

0.05 0.1 0.15 0.2

0 –1 –2 –3 –3 –2 –1

0 X1

Figure 12.1 Example of a multivariate normal distribution. The multivariate random variable X is a vector of two univariate variables, X1 and X2, each with 5000 observations: X = [X1, X2]. The mean vector m = [0,0], as both X1 and X2 have means = 0. The standard deviations of both X1 and X2 = 1, hence the matrix D(s) of sample standard ⎡1 0⎤ deviations = ⎢ ⎥ . (A) Histogram that illustrates the frequency distribution of X1; ⎣0 1⎦ (B) histogram that illustrates the frequency distribution of X2. The correlation between X1 and X2 = 0.75 and is illustrated in the scatterplot (C). ⎡ 1 0.75⎤ Thus, the matrix P of sample correlations = ⎢ ⎥ . The joint multivariate probability ⎣0.75 1 ⎦ distribution of X is shown as a mesh plot in (D). This distribution can also be represented as a contour plot (E), where each contour represents a slice through the mesh plot. Thus, for both X1 and X2 close to 0, the contour label = 0.2 means that the joint probability of obtaining these two values is approximately 0.2. The direction and eccentricity of the contour ellipses (or slices) reflect the correlation between the variables X1 and X2 (C). If X1 and X2 were completely uncorrelated, the contours would be circular. If X1 and X2 were perfectly correlated, the contours would collapse to a single straight line in bivariate space.

Testing for Multivariate Normality

Many multivariate tests assume that the multivariate data or their residuals are multivariate normally distributed. Tests for univariate normality (described in Chap-

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The Multivariate Normal Distribution

ter 11) are used routinely with regression, ANOVA, and other univariate statistical analyses, but tests for multivariate normality are performed rarely. This is not for the lack of availability of such tests. Indeed, more than fifty such tests for multivariate normality have been proposed (reviews in Koizol 1986, Gnanadesikan 1997, and Mecklin and Mundfrom 2003). However, these tests are not included in statistical software packages, and no single test yet has been found to account for the large number of ways that multivariate data can depart from normality (Mecklin and Mundfrom 2003). Because these tests are complex, and often yield conflicting results, they have been called “academic curiosities, seldom used by practicing statisticians” (Horswell 1990, quoted in Mecklin and Mundfrom 2003). A common shortcut for testing for multivariate normality is simply to test whether each individual variable within the multivariate dataset is normally distributed (see Chapter 11). If any of the individual variables is not normally distributed, then it is not possible for the multivariate dataset to be multivariate normally distributed. However, the converse is not true. Each univariate measurement can be normally distributed but the entire dataset still may not be multivariate normally distributed (Looney 1995). Additional tests for multivariate normality should be performed even if each individual variable is found to be univariate normally distributed. One test for multivariate normality is based on measures of multivariate skewness and kurtosis (see Chapter 3). This test was developed by Mardia (1970), and extended by Doornik and Hansen (2008). The computations involved in Doornik and Hansen’s test are comparatively simple and the algorithm is provided completely in their 2008 paper.4 We tested the Darlingtonia data (see Table 12.1) for multivariate normality using Doornik and Hansen’s test. Even though all the individual measurements were normally distributed (P > 0.5, all variables, using the Kolmogorov-Smirnov goodness-of-fit test), the multivariate data departed modestly from multivari4

Doornik and Hansen’s (2008) extension is more accurate (it yields the expected Type I error in simulations) and has better statistical power than Mardia’s test both for small (50 > N > 7) and large (N > 50) sample sizes (Doornik and Hansen 2008; Mecklin and Mundfrom 2003). It is also easier to understand and program than the alternative procedure described by Royston (1983). For a multivariate dataset with m observations and n measurements, Doornik and Hansen’s test statistic En is calculated as En = Z1T Z1 + Z T2 Z2, where Z1 is an n-dimensional column vector of transformed skewnesses of the multivariate data and Z2 is an n-dimensional column vector of transformed kurtoses of the multivariate data. Transformations are given in Doornik and Hansen (2008). The test-statistic En is asymptotically distributed as a χ2 random variable with 2n degrees of freedom. (AME has written an R function to carry out Doornik and Hansen’s test. The code can be downloaded from harvardforest.fas.harvard.edu/ellison/publications/primer/datafiles.)

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ate normality (P = 0.006). This lack of fit was due entirely to the measurement of pitcher keel-width. With this variable removed, the remaining data passed the test for multivariate normality (P = 0.07).

Measurements of Multivariate Distance Many multivariate methods quantify the difference among individual observations, samples, treatment groups, or populations. These differences most frequently are expressed as distances between observations in multivariate space. Before we plunge into the analytical techniques, we describe how distances are calculated between individual observations or between centroids that represent means for entire groups. Using the sample data in Table 12.1, we could ask: How far apart (or different) are two individual plants from one another in the multivariate space created using the morphological variables? Measuring Distances between Two Individuals

Let’s start by considering only two individuals, Plant 1 and Plant 2 of Table 12.1, and two of the variables, pitcher height (Variable 1) and spread of its fishtail appendage (Variable 2). For these two plants, we could measure the morphological distance between them by plotting the points in two dimensions and measuring the shortest distance between them (Figure 12.2). This distance, obtained by applying the Pythagorean theorem,5 is called the Euclidean distance and is calculated as di , j = ( yi ,1 − y j ,1 )2 + ( yi ,2 − y j ,2 )2

(12.15)

In Equation 12.15, the plant is indicated by the subscript letter i or j, and the variable by subscript 1 or 2. To compute the distance between the measurements, we square the difference in heights, add to this the square of the difference in spreads, and take the square root of the sum. The result for these

Pythagoras

Pythagoras of Samos (ca. 569–475 B.C.) may have been the first “pure” mathematician. He is remembered best for being the first to offer a formal proof of what is now known as the Pythagorean Theorem: the sum of the squares of the lengths of the two legs of a right triangle equals the square of the length of its hypotenuse: a2 + b2 = c2. Note that the Euclidean distance we are using is the length of the hypotenuse of an implicit right triangle (see Figure 12.2). Pythagoras also knew that the Earth was spherical, although by placing it at the center of the universe he misplaced it a few billion light-years away from its actual position.

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Spread of the fishtail appendage (mm)

Plant j (yj,1, yj,2)

59 di,j =√(yi,1 – yj,1)2 + (yi,2 – yj,2)2

58 57 56 55

Figure 12.2 In two dimensions, the Euclidean distance di,j is the straight-line distance between the points. In this example, the two morphological variables that we measured are plant height and spread of the fishtail appendage of the carnivorous cobra lily, Darlingtonia californica (see Table 12.1). Here the measurements for two individual plants are plotted in bivariate space. Equation 12.15 is used to calculate the Euclidean distance between them.

Plant i (yi,1, yi,2) 400

450

500 550 Height (mm)

600

650

two plants is a distance di,j = 241.05 mm. Because the original variables were measured in millimeters, when we subtract one measure from another, our result is still in millimeters. Applying subsequent operations of squaring, summing and taking the square root brings us back to a distance measurement that is still in units of millimeters. However, the units of the distance measurements will not stay the same unless all of the original variables were measured in the same units. Similarly, we can calculate the Euclidean distance between these two plants if they are described by three variables: height, spread, and mouth diameter (Figure 12.3). We simply add another squared difference—here the difference between mouth diameters—to Equation 12.15: di, j = ( yi,1 − y j,1 )2 + ( yi,2 − y j,2 )2 + ( yi,3 − y j,3 )2

(12.16)

Pythagoras founded a scientific-cum-religious society in Croton, in what is now southern Italy. The inner circle of this society (which included both men and women) were the mathematikoi, communitarians who renounced personal possessions and were strict vegetarians. Among their core beliefs were that reality is mathematical in nature; that philosophy is the basis of spiritual purification; that the soul can obtain union with the divine; and that some symbols have mystical significance. All members of the order were sworn to the utmost loyalty and secrecy, so there is scant biographical data about Pythagoras.

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Plant i (yi,1, yi,2, yi,3) Mouth diameter (mm)

400

35 30

di,j = √(yi,1 – yj,1)2 + (yi,2 – yj,2)2+ (yi,3 – yj,3)2

25 Plant j (yj,1, yj,2, yj,3)

20 59

ail ht ) fis m of e (m ad g re da Sp pen ap

56 55

420

540 ) 480 (mm t h g i e H

600

Figure 12.3 Measuring Euclidean distance in three-dimensional space. Mouth diameter is added to the two morphological variables shown in Figure 12.2, and the three variables are plotted in three-dimensional space. Equation 12.16 is used to calculate Euclidean distance, which is 241.58. Notice that this distance is virtually identical to the Euclidean distance measured in two dimensions (241.05; see Figure 12.2). The reason is that the third variable (mouth diameter) has a much smaller mean and variance than the first two variables and so it does not affect the distance measurement very much. For this reason, variables should be standardized (using Equation 12.17) before calculating distances between individuals.

The result of applying Equation 12.16 to the two plants is a morphological distance of 241.58 mm—a mere 0.2% change from the distance obtained using only the two variables height and spread. Why are these two Euclidean distances not very different? Because the magnitude of plant height (hundreds of millimeters) is much greater than the magnitude of either spread or mouth diameter (tens of millimeters), the measurement of plant height dominates the distance calculations. In practice, therefore, it is essential to standardize the variables prior to computing distances. A convenient standardization is to transform each variable by subtracting its sample mean from the value of each observation for that variable, and then dividing this difference by the sample standard deviation: Z=

(Yi − Y ) s

(12.17)

The result of this transformation is called a Z-score. The Z-score controls for differences in the variance of each of the measured variables, and can also be

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TABLE 12.3 Standardization of multivariate data Site

TJH TJH ...

Plant Height Mouth Tube

1 2 ...

Keel Wing1 Wing2 Wspread Hoodmass Tubemass Wingmass

0.381 1.202 –1.062 0.001 0.516 0.156 –2.014 –1.388 –0.878 –0.228 –0.802 –1.984 ...

...

–1.035 –0.893 ...

1.568 –0.881 ...

0.602 –1.281 ...

0.159 –0.276 ...

The original data are measurements of 7 morphological variables and 3 biomass variables on 89 individuals of Darlingtonia californica collected at four sites (see Table 12.1). The first two rows of the data are illustrated after standardization. Standardized values are calculated by subtracting the sample mean of each variable from each observation and dividing this difference by the sample standard deviation (see Equation 12.17).

used to compare measurements that are not in the same units. The standardized values (Z-scores) for Table 12.1 are given in Table 12.3. The distance between these two plants for the two standardized variables height and spread is 2.527, and for the three standardized variables height, spread, and mouth diameter is 2.533. The absolute difference between these two distances is a modest 1%. However, this difference is five times greater than the difference between the distances calculated for the non-standardized data. When the data have been transformed with Z-scores, what are the units of the distances? If we were working with our original variables, the units would be in millimeters. But standardized variables are always dimensionless; Equation 12.17 gives us units of mm × mm–1, which cancel each other out. Normally, we think of Z-scores as being in units of “standard deviations”—how many standard deviations a measurement is from the mean. For example, a plant with a standardized height measurement of 0.381 is 0.381 standard deviations larger than the average plant. Although we can’t draw four or more axes on a flat piece of paper, we can still calculate Euclidean distances between two individuals if they are described by n variables. The general formula for the Euclidean distance based on a set of n variables is: d i, j =

∑ ( yi,k − y j,k )2

(12.18)

k =1

Based on all seven standardized morphological variables, the Euclidean distance between the two plants in Table 12.3 is 4.346.

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Measuring Distances between Two Groups

Typically, we want to measure distances between samples or experimental treatment groups, not just distances between individuals. We can extend our formula for Euclidean distance to measure the Euclidean distance between means of any arbitrary number of groups g: g

d i, j =

∑ (Yi,k − Y j,k )2

(12.19)

k =1

– where Y i,k is the mean of variable i in group k. The Yi in Equation 12.19 can be univariate or multivariate. Again, so that no one variable dominates the distance calculation, we usually standardize the data (see Equation 12.17) before calculating means and the distances between groups. From Equation 12.19, the Euclidean distance between the populations at DG and TJH based on all seven morphological variables is 1.492 standard deviations. Other Measurements of Distance

Euclidean distance is the most commonly used distance measure. However, it is not always the best choice for measuring the distance between multivariate objects. For example, a common question in community ecology is how different two sites are based on the occurrence or abundance of species found at the two sites. A curious, counterintuitive paradox is that two sites with no species in common may have a smaller Euclidean distance than two sites that share at least some species! This paradox is illustrated using hypothetical data (Orlóci 1978) for three sites x1, x2, and x3, and three species y1, y2, y3. Table 12.4 gives the site × species matrix, and Table 12.5 gives all the pairwise Euclidean

This matrix illustrates the paradox of using Euclidean distances for measuring similarity between sites based on species abundances. Each row represents a site, and each column a species. The values are the number of individuals of each species yi at site xj. The paradox is that the Euclidean distance between two sites with no species in common (such as sites x1 and x2) may be smaller than the Euclidean distance between two sites that share at least some species (such as sites x1 and x3).

TABLE 12.4 Site × species matrix Species Site

x1 x2 x3

0 1 0

1 0 4

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TABLE 12.5 Pair-wise Euclidean distances between sites Site Site

x1 x2 x3

0 1.732 4.243

1.732 0 5.745

4.243 5.745 0

403

This matrix is based on the species abundances given in Table 12.4. Each entry of the matrix is the Euclidean distance between site xj and xk. Note that the distance matrix is symmetric (see Appendix); for example, the distance between site x1 and x2 is the same as the distance between site x2 and x1. The diagonal elements are all equal to 0, because the distance between a site and itself = 0. This matrix of distance calculations demonstrates that Euclidean distances may give counterintuitive results if the data include many zeros. Sites x1 and x2 share no species in common, but their Euclidean distance is smaller than the Euclidean distance between sites x1 and x3, which share the same set of species.

distances between the sites. In this simple example, sites x1 and x2 have no species in common, and the Euclidean distance between them is 1.732 species (see Equation 12.14): dx1, x2 = (1 − 0)2 + (1 − 0)2 + (1 − 0)2 = 1.732 In contrast, sites x1 and x3 have all their species in common (both y2 and y3 occur at these two sites), but the distance between them is 4.243 species: dx1, x3 = (0 − 0)2 + (1 − 4)2 + (1 − 4)2 = 4.243 Ecologists, environmental scientists, and statisticians have developed many other measures to quantify the distance between two multivariate samples or populations. A subset of these are given in Table 12.6; Legendre and Legendre (1998) and Podani and Miklós (2002) discuss many more. These distance measurements fall into two categories: metric distances and semi-metric distances. Metric distances have four properties: 1. The minimum distance = 0, and if two objects (or samples) x1 and x2 are identical, then the distance d between them also equals 0: x1 = x2 ⇒ d(x1,x2) = 0. 2. The distance measurement d is always positive if two objects x1 and x2 are not identical: x1 ≠ x2 ⇒ d(x1,x2) > 0. 3. The distance measurement d is symmetric: d(x1,x2) = d(x2,x1). 4. The distance measurement d satisfies the triangle inequality: for three objects x1, x2 and x3, d(x1,x2) + d(x2,x3) ≥ d(x1,x3).

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TABLE 12.6 Some common measures of distance or dissimilarity used by ecologists Name

Formula

Euclidean

d i, j =

Property n

∑ ( yi,k − yj,k )2

Metric

k =1

Manhattan (aka City Block)

Chord

Mahalanobis

di , j = ∑ y i ,k − y j ,k

⎛ ⎜ ⎜ d i, j = 2 × ⎜1 − ⎜ ⎜ ⎝

∑ yi,k y j,k k =1

∑ ∑ k =1

yi2,k

k =1

y 2j,k

⎞ ⎟ ⎟ ⎟ ⎟ ⎟ ⎠

Metric

d y i , yj = d i, j V −1d Ti, j V=

Chi-square

Metric

k =1

Metric

1 [(mi − 1)C i + (m j − 1)C j ] mi + m j − 2

⎡ ⎛ ⎢ ⎜ n ⎢ m m y jk y 1 di , j = ∑ ∑ y ij × ∑ ⎢ n × ⎜ n ik − n ⎜ k =1 ⎢ i =1 j =1 ⎢ ∑ y jk ⎜ ∑ yik ∑ yjk ⎝ k =1 k =1 ⎢⎣ k =1

⎞ ⎟ ⎟ ⎟ ⎟ ⎠

2⎤

⎥ ⎥ ⎥ ⎥ ⎥ ⎥⎦

Metric

Bray-Curtis

d i, j =

∑ yi,k − y j,k k =1 n

Semi-metric

∑ ( yi,k + y j,k ) k =1

Jaccard

d i, j =

a +b a +b +c

Metric

Sørensen’s

d i, j =

a +b a + b + 2c

Semi-metric

The Euclidean, Manhattan (or City Block), Chord, and Bray-Curtis distances are used principally for continuous numerical data. Jaccard and Sørensen’s distances are used for measuring distances between two samples that are described by presence/absence data. In Jaccard and Sørensen’s distances, a is the number of objects (e.g., species) that occur only in yi, b is the number of objects that occur only in yj, and c is the number of objects that occur in both yi and yj. The Mahalanobis distance applies only to groups of samples yi and yj, each of which contains respectively mi and mj samples. In the equation for Mahalanobis distance, d is the vector of differences between the means of the m samples in each group, and V is the pooled within-group sample variance-covariance matrix, calculated as shown, where Ci is the sample variance-covariance matrix for yi (see Equation 12.3).

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Euclidean, Manhattan, Chord, Mahalanobis, chi-square, and Jaccard distances6 are all examples of metric distances.

6 If you’ve used the Jaccard index of similarity before, you may wonder why we refer to it in Table 12.6 as a distance or dissimilarity measure. Jaccard’s (1901) coefficient was developed to describe how similar two communities are in terms of shared species si,j c si , j = a+b+c where a is the number of species that occur only in Community i, b is the number of species that occur only in Community j, and c is the number of species they have in common. Because measures of similarity take on their maximum value when objects are most similar, and measures of dissimilarity (or distance) take on their maximum value when objects are most different, any measure of similarity can be transformed into a measure of dissimilarity or distance. If a measure of similarity s ranges from 0 to 1 (as does Jaccard’s coefficient), it can be transformed into a measure of distance d by using one of the following three equations:

d = 1 − s, d = 1 − s , or d = 1 − s 2

Thus, in Table 12.6, the Jaccard distance equals 1 – the Jaccard coefficient of similarity. The reverse transformation (e.g., s = 1 – d) can be used to transform measures of distance into measures of similarity. If the distance measurement is not bounded (i.e., it ranges from 0 to ∞), it must be normalized to range between 0 and 1: dnorm =

d d − dmin or dnorm = dmax dmax − dmin

Although these algebraic properties of similarity indices are straightforward, their statistical properties are not. Similarity indices as used in biogeography and community ecology are very sensitive to variation in sample size (Wolda 1981; Jackson et al. 1989). In particular, small communities may have relatively high similarity coefficients even in the absence of any unusual biotic forces because their faunas are dominated by a handful of common, widespread species. Rare species are found mostly in larger faunas, and they will tend to reduce the similarity index for two communities even if the rare species also occur at random. Similarity indices and other biotic indices should be compared to an appropriate null model that controls for variation in sample size or the number of species present (Gotelli and Graves 1996). Either Monte Carlo simulations or probability calculations can be used to determine the expected value of a biodiversity or similarity index at small sample sizes (Colwell and Coddington 1994; Gotelli and Colwell 2001). The measured similarity in species composition of two assemblages depends not only on the number of species they share, but the dispersal potential of the species (most simple models assume species occurrences are equiprobable), and the composition and size of the source pool (Connor and Simberloff 1978; Rice and Belland 1982). Chao et al. (2005) introduce an explicit sampling model for the Jaccard index that accounts not only for rare species, but also for species that are shared but were undetected in any of the samples.

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Semi-metric distances satisfy only the first three of these properties, but may violate the triangle inequality. Bray-Curtis and Sørensen’s measures are semimetric distances. A third type of distance measure, not used by ecologists, is nonmetric, which violates the second property of metric distances and may take on negative values.

Ordination Ordination techniques are used to order (or ordinate) multivariate data. Ordination creates new variables (called principal axes) along which samples are scored or ordered. This ordering may represent a useful simplification of patterns in complex multivariate data sets. Used in this way, ordination is a datareduction technique: beginning with a set of n variables, the ordination generates a smaller number of variables that still illustrate the important patterns in the data. Ordination also can be used to discriminate or separate samples along the axis. Ecologists and environmental scientists routinely use five different types of ordination—principal component analysis, factor analysis, correspondence analysis, principal coordinates analysis, and non-metric multidimensional scaling—each of which we discuss in turn. We discuss in depth the details of principal component analysis because the concepts and methods are similar to those used in other ordination techniques. Our discussion of the other four ordination methods is somewhat briefer. Legendre and Legendre (1998) is a good ecological guide to the details of basic multivariate analysis. Principal Component Analysis

Principal component analysis (PCA) is the most straightforward way to ordinate data. The idea of PCA is generally credited to Karl Pearson (1901),7 of Pearson chi-square and correlation fame, but Harold Hotelling 8 (of the Hotelling T 2 test) developed the computational methods in 1933. The primary use of PCA is to reduce the dimensionality of multivariate data. In other words, we use PCA to create a few key variables (each of which is a composite of many of our original variables) that characterize as fully as possible the variation in a multivariate dataset. The most important attribute of PCA is that the new variables are not correlated with one another. These uncorrelated variables can be used in multiple regression (see Chap-

Pearson’s goal in his 1901 paper was to assign individuals to racial categories based on multiple biometric measurements. See Footnote 3 in Chapter 11 for a biographical sketch of Karl Pearson.

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ter 9) or ANOVA (see Chapter 10) without fear of multicollinearity. If the original variables are normally distributed, or have been transformed (see Chapter 8) or standardized (see Equation 12.17) prior to analysis, the new variables resulting from the PCA also will be normally distributed, satisfying one of the key requirements of parametric tests that are used for hypothesis testing. PCA: THE CONCEPT

Figure 12.4 illustrates the basic concept of a principal component analysis. Imagine you have counted the number of individuals of two species of grassland sparrows in each of ten prairie plots. These data can be plotted in a graph in which the abundance of Species A is on the x-axis and the abundance of Species B is on the y-axis. Each point in the graph represents the data from a different plot. We now create a new variable, or first principal axis,9 that passes through the center of the cloud of the data points. Next, we calculate the value for each of the ten plots along this new axis, and then graph them on the axis. For each point, this calculation uses the numbers of both Species A and Species B to generate a single new value, which is the principal component score on the first principal axis. Although the original multivariate data had two observations for each plot, the principal component score reduces these two observations to a single number. We have effectively reduced the dimensionality of the data from two dimensions (Species A abundance and Species B abundance) to one dimension (first principal axis). In general, if you have measured j = 1 to n variables Yj for each replicate, you can generate n new variables Zj that are all uncorrelated with one another (all the off-diagonal elements in P = 0). We do this for two reasons. First, because the Zj’s are uncorrelated, they each can be thought of as measuring a different

Harold Hotelling

Harold Hotelling (1895–1973) made significant contributions to statistics and economics, two fields from which ecologists have pilfered many ideas. His 1931 paper extending the t-test to the multivariate case also introduced confidence intervals to statisticians, and his 1933 paper developed the mathematics of principal components. As an economist, he was best known for championing approaches using marginal costs and benefits—the neoclassical tradition in economics based on Vilfredo Pareto’s Manual of Political Economy. This tradition also forms the basis for many optimization models of ecological and evolutionary trade-offs.

9 The term “principal axis” is derived from optics, where it is used to refer to the line (also known as the optical axis) that passes through the center of curvature of a lens so that a ray of light passing along that line is neither reflected nor refracted. It is also used in physics to refer to an axis of symmetry, such that an object will rotate at a constant angular velocity about its principal axis without applying additional torque.

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(A) Abundance of Species B

408

Abundance of Species A

(B)

First principal axis

Figure 12.4 Construction of a principal component axis. (A) For a hypothetical set of 10 prairie plots, the number of individuals of two species of grassland sparrows are measured in each plot. Each plot can be represented as a point in a bivariate scatterplot with the abundance of each species graphed on the two axes. The first principal axis (blue line) passes through the major axis of variation in the data. The second principal axis (short black line) is orthogonal (perpendicular) to the first, and accounts for the small amount of residual variation not already incorporated in the first principal axis. (B) The first principal axis is then rotated, becoming the new axis along which the plot scores are ordinated (ordered). The vertical black line indicates the crossing of the second principal axis. Points to the right of the vertical line have positive scores on the first principal axis, and points to the left of the vertical line have negative scores. Although the original multivariate data had two measurements for each plot, most of the variation in the data is captured by a single score on the first principal component, allowing the plots to be ordinated along this axis.

and independent “dimension” of the multivariate data. Note that these new variables are not the same as the Z-scores described earlier (see Equation 12.17). Second, the new variables can be ordered based on the amount of variation they explain in the original data. Z1 has the largest amount of variation in the data (it is called the major axis), Z2 has the next-largest amount of variation in the data, and so on (var(Z1) ≥ var(Z2) ≥ … ≥ var(Zn)). In a typical ecological data set, most of the variation in the data is captured in the first few Zj ’s, and we can discard the subsequent Zj ’s that account for the residual variation. Ordered this way, the Zj’s are called principal components. If PCA is informative, it reduces a large number of original, correlated variables to a small number of new, uncorrelated variables.

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Principal component analysis is successful to the extent that there are strong intercorrelations in the original data. If all of the variables are uncorrelated to begin with, we don’t gain anything from the PCA because we cannot capture the variation in a handful of new, transformed variables; we might as well base the analysis on the untransformed variables themselves. Moreover, PCA may not be successful if it is used on unstandardized variables. Variables need to be standardized to the same relative scale, using transformations such as Equation 12.17, so that the axes of the PCA are not dominated by one or two variables that have large units of measurement. So how does it work? By way of example, let’s examine some of the results of a PCA applied to the Darlingtonia data in Table 12.1. For this analysis, we used all ten variables—height, mouth diameter, tube and keel diameters, lengths and spread of the wings of the fishtail appendage, and masses of the hood, tube, and appendage—which we will call Y1 through Y10. The first principal component, Z1, is a new variable (don’t panic, details will follow), whose value for each observation is AN EXAMPLE OF A PCA

Z1 = 0.31Y1 + 0.40Y2 – 0.002Y3 – 0.18Y4 + 0.39Y5 + 0.37Y6 + 0.26Y7 + 0.40Y8 + 0.38Y9 + 0.23Y10

(12.20)

We generate the first principal component score (Z1) for each individual plant by multiplying each measured variable times the corresponding coefficient (the loading) and summing the results. Thus, we take 0.31 times plant height, plus 0.40 times plant mouth diameter, and so on. For the first replicate in Table 12.1, Z1 = 1.43. Because the coefficients for some of the Yj’s may be negative, the principal component score for a particular replicate may be positive or negative. The variance of Z1 = 4.49 and it accounts for 46% of the total variance in the data. The second and third principal components have variances of 1.63 and 1.46, and all others are < 0.7. Where did Equation 12.20 come from? Notice first that Z1 is a linear combination of the ten Y variables. As we described above, we have multiplied each variable Yj by a coefficient aij , also known as the loading, and added up all these products to give us Z1. Each additional principal component is another linear combination of all of the Yj ’s: Zj = ai1Y1 + ai2Y2 + … + ainYn

(12.21)

The aij ’s are the coefficients for factor i that are multiplied by the measured value for variable j. Each principal component accounts for as much variance in the data as possible, subject to the condition that all the Zj’s are uncorrelated. This

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condition ensures that the Zj’s are independent and orthogonal. Therefore, when we plot the Zj’s, we are seeing relationships (if there are any) between independent variables. The calculation of both the coefficients aij and their associated variance is relatively simple. We start by standardizing our data by applying Equation 12.17, and then computing the sample variancecovariance matrix C (see Equation 12.3) of the standardized data. Note that C computed using standardized data is the same as the correlation matrix P (see Equation 12.5) computed using the raw data. We then calculate the eigenvalues λ1 … λn of the sample variance-covariance matrix and their associated eigenvectors aj (see the Appendix for methods). The jth eigenvalue is the variance of Zj, and the loadings aij are the elements of the eigenvectors. The sum of all the eigenvalues is the total variance explained: CALCULATING PRINCIPAL COMPONENTS

vartotal = ∑ λ j j =1

and the proportion of variance explained by each component Zj is varj =

lj n

∑lj j =1

If we multiply varj by 100, we get the percent of variance explained. Because there are as many principal components as there are original variables, the total amount of variance explained by all the principal components = 100%. The results of these calculations for the Darlingtonia data in Table 12.1 are summarized in Tables 12.7 and 12.8. Because PCA is a method for simplifying multivariate data, we are interested in retaining only those components that explain the bulk of the variation in the data. There is no absolute cutoff point at which we discard principal components, but a useful graphical tool to examine the contribution of each principal component to the overall PCA is a scree plot (Figure 12.5). This plot illustrates the percent of variance (derived from the eigenvalue) explained by each component in decreasing order. The scree plot resembles the profile of a mountainside down which a lot of rubble has fallen (known as a scree slope). The data used to construct the scree plot are tabulated in Table 12.7. In a scree plot, we look for a sharp bend or change in slope of the ordered eigenvalues. We retain those components that contribute to the mountainside, and ignore the rubble at the bottom. In this example, Components 1–3 appear

HOW MANY COMPONENTS TO USE?

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TABLE 12.7 Eigenvalues from principal component analysis Principal component

Eigenvalue li

Proportion of variance explained

Z1 Z2 Z3 Z4 Z5 Z6 Z7 Z8 Z9 Z10

4.51 1.65 1.45 0.73 0.48 0.35 0.25 0.22 0.14 0.05

0.458 0.167 0.148 0.074 0.049 0.036 0.024 0.023 0.014 0.005

Cumulative proportion of variance explained

0.458 0.625 0.773 0.847 0.896 0.932 0.958 0.981 0.995 1.000

These eigenvalues were obtained from analysis of the standardized measurements of 7 morphological variables and 3 biomass variables from 89 individuals of the cobra lily Darlingtonia californica collected at 4 sites (see Table 12.3). In a principal component analysis, the eigenvalue measures the proportion of variance in the original data explained by each principal component. The proportion of variance explained and the cumulative proportion explained are calculated from the sum of the eigenvalues (∑λj = 9.83). The proportion of variance explained is used to select a small number of principal components that capture most of the variation in the data. In this data set, the first 3 principal components account for 77% of the variance in the original 10 variables.

TABLE 12.8 Eigenvectors from a principal component analysis ⎡ 0.31 ⎤ ⎢ ⎥ ⎢ 0.40 ⎥ ⎢−0.002⎥ ⎢ ⎥ ⎢−0.18 ⎥ ⎢ 0.39 ⎥ ⎥ a1 = ⎢ ⎢ 0.37 ⎥ ⎢ 0.26 ⎥ ⎢ ⎥ ⎢ 0.40 ⎥ ⎢ ⎥ ⎢ 0.38 ⎥ ⎢⎣ 0.23 ⎥⎦

⎡ −0.42⎤ ⎢ ⎥ ⎢ −.025⎥ ⎢ −0.10⎥ ⎢ ⎥ ⎢−0.07 ⎥ ⎢ 0.28⎥ ⎥ a2 = ⎢ ⎢ 0.37⎥ ⎢ 0.43⎥ ⎢ ⎥ ⎢ −0.18⎥ ⎢ ⎥ ⎢ −0.41⎥ ⎢⎣ 0.35⎥⎦

⎡ 0.17 ⎤ ⎢ ⎥ ⎢ −0.11 ⎥ ⎢ −.74 ⎥ ⎢ ⎥ ⎢ −.58 ⎥ ⎢ −0.004 ⎥ ⎥ a3 = ⎢ ⎢ 0.09 ⎥ ⎢ 0.19 ⎥ ⎢ ⎥ ⎢ −0.08 ⎥ ⎢ ⎥ ⎢ 0.04 ⎥ ⎢⎣ 0.11 ⎥⎦

The first three eigenvectors from a principal component analysis of the measurements in Table 12.1. Each element in the eigenvector is the coefficient or loading that is multiplied by the value of the corresponding standardized variable (see Table 12.3). The products of the loadings and the standardized measurements are summed to give the principal component score. Thus, using the first eigenvector, 0.31 is multiplied by the first measurement (plant height), and this is added to 0.40 multiplied by the second measurement (plant mouth diameter), and so on. In matrix notation, we say that the principal component scores Zj for each observation y, which consists of ten measurements y1 through y10, are obtained by multiplying ai by y (see Equations 12.20–12.23).

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0.458 4

3 Variance

412

0.625

0.773

0.847 0.896 0.932

0.958 0.981 0.995 1.000

0 1

Component

Figure 12.5 A scree plot for principal component analysis of the standardized data in Table 12.3. In a scree plot, the percent of variance (the eigenvalue) explained by each component (see Table 12.7) is shown in decreasing order. The scree plot resembles the profile of a mountainside down which a lot of rubble has fallen (known as a scree slope). In this scree plot, Component 1 clearly dominates the mountainside, although components 2 and 3 are also informative. The remainder is rubble.

useful, in total explaining 77% of the variance, whereas Components 4–10 look like rubble, none of them explaining more than 7% of the remaining variance. Jackson (1993) discusses a wide variety of heuristic methods (such as the scree plot) and statistical methods for choosing how many principal components to use. His results suggest that scree plots tend to overestimate by one the number of statistically meaningful components. Now that we have selected a small number of components, we want to examine them in more detail. Component 1 was described already, in Equation 12.20:

WHAT DO THE COMPONENTS MEAN?

Z1 = 0.31Y1 + 0.40Y2 – 0.002Y3 – 0.18Y4 + 0.39Y5 + 0.37Y6 + 0.26Y7 + 0.40Y8 + 0.38Y9 + 0.23Y10

This component is the matrix product of the first eigenvector a1 (see Table 12.8) and the matrix of multivariate observations Y that had been standardized using Equation 12.17. Most of the loadings (or coefficients) are positive, and all are of approximately the same magnitude except the two loadings for variables relat-

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ed to the leaf tube (tube diameter Y3 and keel diameter Y4). Thus, this first component Z1 seems to be a good measure of pitcher “size”—plants with tall pitchers, large mouths, and large appendages will all have larger Z1 values. Similarly, Component 2 is the product of the second eigenvector a2 (see Table 12.8) and the matrix of standardized observations Y: Z2 = –0.42Y1 – 0.25Y2 – 0.10Y3 –0.07Y4 + 0.28Y5 + 0.37Y6 + 0.43Y7 – 0.18Y8 – 0.41Y9 + 0.35Y10

(12.22)

For this component, loadings on the six variables related to pitcher height and diameter are negative, whereas loadings on the four variables related to the fishtail appendage are positive. Because all our variables have been standardized, relatively short, skinny pitchers will have negative height and diameter values, whereas relatively tall, stout pitchers will have positive height and diameter values. The value of Z2 consequently reflects trade-offs in shape. Short plants with large appendages will have large values of Z2, whereas tall plants with small appendages will have small values of Z2. We can think of Z2 as a variable that describes pitcher “shape.” We generate the second principal component score for each individual plant by again substituting its observed values y1 through y10 into Equation 12.22. Finally, Component 3 is the product of the third eigenvector a3 (see Table 12.8) and the matrix of standardized observations Y: Z3 = 0.17Y1 – 0.11Y2 + 0.74Y3 + 0.58Y4 – 0.004Y5 + 0.09Y6 + 0.19Y7 – 0.08Y8 + 0.04Y9 + 0.11Y10

(12.23)

This component is overwhelmingly dominated by large positive coefficients for tube (Y3) and keel (Y4) diameters—measurements of structures that support the pitcher. Z3 will be large for “fat” plants and small for “skinny” plants. Because insects are trapped within the tube, plants with large Z3 scores may be able to trap larger prey than plants with smaller Z3 scores. ARE ALL THE LOADINGS MEANINGFUL? In creating the principal component scores, we used the loadings from all of the original variables. But some loadings are large and others are close to zero. Should we use them all, or only those that are above some cutoff value? There is little agreement on this point (Peres-Neto et al. 2003), and if PCA is used only for exploratory data analysis, it probably doesn’t matter much whether or not small loadings are retained. But if you use principal component scores for hypothesis testing, you must explicitly state which loadings were used, and how you decided to include or exclude them. If you are testing for interactions among principal axes, it is important to retain all of them.

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Finally, we may want to examine differences among the four sites in their principal component scores. We can illustrate these differences by plotting the principal component scores for each plant on two axes, and coding the points (= replicates) for each group using different colors or symbols (Figure 12.6). In this plot, we illustrate the scores for the first two principal components, Z1 and Z2. We could construct similar plots for the other meaningful principal components (e.g., Z1 versus Z3; Z2 versus Z3). Figure 12.6 was generated using all the loadings in Equations 12.20 and 12.22. An important feature of PCA is that the positions of the principal component scores for each replicate (such as the points in Figure 12.6) have the same Euclidean distance between them as do the original data in multivariate space. This property holds only when the Euclidean distances are calculated using all the principal components (Zj’s). If you calculate Euclidean distances using only the first few principal components, they will not be equal to the distances between the original data points. If the data originally were collected along with

USING THE COMPONENTS TO TEST HYPOTHESES

High Divide (HD) L.E. Hunter’s Fen (LEH) Day’s Gulch (DG) T.J. Howell’s Fen (TJH)

Principal component 2

414

3 1 –1 –3 –5 –4

–2

0 2 4 Principal component 1

Figure 12.6 Plot illustrating the first two principal component scores from a PCA on the data in Table 12.3. Each point represents an individual plant, and the colors represent the different sites in the Siskiyou Mountains of southern Oregon. Although there is considerable overlap among the groups, there is also some distinct separation: the TJH samples (light blue) have lower scores on principal component 2, and the HD samples (black) have lower scores on the first principal component. Univariate or multivariate tests can be used to compare principal component scores among the populations (see Figure 12.7).

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spatial information (such as x,y or latitude-longitude coordinates where the data were collected), the principal component scores could be plotted against their spatial coordinates. Such plots can provide information on multivariate spatial relationships. We can treat the principal component scores as a simple univariate response variable and use ANOVA (see Chapter 10) to test for differences among treatment groups or sample populations. For example, an ANOVA on the first principal component score of the Darlingtonia data gives the result that there are significant differences among sites (F3,79 = 9.26, P = 2 × 10–5), and that all pair-wise differences among sites, except for the DG–LEH comparison, also are significantly different (see Chapter 10 for details on calculating a posteriori comparisons). We conclude that plant size varies systematically across sites (Figure 12.7). We obtain similar results for plant shape (F3,79 = 5.36, P = 0.002), but only three of the pairwise comparisons—DG versus TJH; HD versus TJH; and LEH versus TJH—are significantly different. This result suggests that plants at TJH have a different shape from plants at all the other sites. The concentration of light blue points (the principal component scores for the plants at TJH) in Figure 12.6 reveals that these plants have lower scores for the second principal component—that is, they are relatively tall plants with small fishtail appendages. Factor Analysis

Factor analysis10 and PCA have a similar goal: the reduction of many variables to few variables. Whereas PCA creates new variables as linear combinations of the original variables (see Equation 12.21), factor analysis considers each of

10 Factor analysis was developed by Charles Spearman (1863–1945). In his 1904 paper, Spearman used factor analysis to measure general intelligence from multiple test scores. The factor model (see Equation 12.24) was used to partition the results from a battery of “intelligence tests” (the Yj’s) into factors (the Fj’s) that measured general intelligence from factors specific to each test (the ej’s). This dubious discovery provided the theoretical underpinnings for Binet’s general tests of intelligence (Intelligence Quotient, or I.Q., Charles Spearman tests). Spearman’s theories were used to develop the British “PlusElevens” examinations, which were administered to schoolboys at age 11 in order to determine whether they would be able to attend a University or a technical/vocational school. Standardized testing at all levels (e.g., MCAT, SAT, and GRE in the United States) is the legacy of Spearman’s and Binet’s work. Stephen Jay Gould’s The Mismeasure of Man (1981) describes the unsavory history of IQ testing and the use of biological measurements in the service of social oppression.

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416

3 1 –1 –3 –5 DG

LEH

TJH

Site

Figure 12.7 Box-plots of the scores of the first principal component from the PCA on the data in Table 12.3. Each box plot (see Figure 3.6) represents one of the four sites specified in Figure 12.6. The horizontal line indicates the sample median, and the box encompasses 50% of the data, from the 25th to the 75th percentile. The “whiskers” encompass 90% of the data, from the 10th to the 90th percentile. Four extreme data points (beyond the 5th and 95th percentiles) are shown as horizontal bars for the LEH site. Scores on Principal Component 1 varied significantly among sites (F3,79 = 9.26, P = 2 × 10–5), and were lowest (and least variable) at HD (see Figure 12.6).

the original variables to be a linear combination of some underlying “factors.” You can think of this as PCA in reverse: Yj = ai1F1 + ai2F2 + … + ainFn + ej

(12.24)

To use this equation, the variables Yj must be standardized (mean = 0, variance = 1) using Equation 12.17. Each aij is a factor loading, the F ’s are called common factors (which each have mean = 0 and variance = 1), and the ej’s are factors specific to the jth variable, also uncorrelated with any Fj , and each with mean = 0. Being a “PCA-in-reverse,” factor analysis usually begins with a PCA and uses the meaningful components as the initial factors. Equation 12.21 gave us the formula for generating principal components: Zj = ai1Y1 + ai2Y2 + … + ainYn

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Because the transformation from Y to Z is orthogonal, there is a set of coefficients a*ij such that Yj = a*i1 Z1 + a *i2 Z2 + … + a *in Zn

(12.25)

For a factor analysis, we keep only the first m components, such as those determined to be important in the PCA using a scree plot: Yj = a *i1 Z1 + a *i2 Z2 + … + a *im Zm + ej

(12.26)

To transform the Zj’s into factors (which are standardized with variance = 1), we divide each of them by its standard deviation, l i , the square root of its corresponding eigenvalue. This gives us a factor model: Yj = bi1F1 + bi2F2 + … + bimFm + ej

(12.27)

where F j = Z j / λ j and bij = a ij* l j . ROTATING THE FACTORS Unlike the Zj’s generated by PCA, the factors Fj in Equation 12.27 are not unique; more than one set of coefficients will solve Equation 12.27. We can generate new factors F *j that are linear combinations of the original factors:

F *j = di1F1 + di2F2 + … + dimF m

(12.28)

These new factors also explain as much of the variance in the data as the original factors. After generating factors using Equations 12.25–12.27, we identify the values for the coefficients dij in Equation 12.28 that give us the factors that are the easiest to interpret. This identification process is called rotating the factors. There are two kinds of factor rotations: orthogonal rotation results in new factors that are uncorrelated with one another, whereas oblique rotation results in new factors that may be correlated with one another. The best rotation is one in which the factor coefficients dij are either very small (in which case the associated factor is unimportant) or very large (in which case the associated factor is very important). The most common type of orthogonal factor rotation is called varimax rotation, which maximizes the sum of the variance of the factor coefficients in Equation 12.28: n

∑ dij2 )

varmax = max imum of var(

j =1

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Factor scores for each observation are computed as the values of the Fj’s for each Yi. Because the original factor scores Fj are linear combinations of the data (working backward from Equation 12.27 to Equation 12.25), we can re-write Equation 12.28 in matrix notation as: COMPUTING AND USING FACTOR SCORES

F* = (DTD)–1DTY

(12.29)

where D is the n × n matrix of factor coefficients dij , Y is the data matrix, and F* is the matrix of rotated factor scores. Solving this equation directly gives us the factor scores for each replicate. Be careful when carrying out hypotheses tests on factor scores. Although they can be useful for describing multivariate data, they are not unique (as there are infinitely many choices of Fj), and the type of rotation used is arbitrary. Factor analysis is recommended only for exploratory data analysis. A BRIEF EXAMPLE We conducted a factor analysis on the same data used for the PCA: 10 standardized variables measured on 89 Darlingtonia plants at four sites in Oregon and California (see Table 12.3). The analysis used four factors, which respectively accounted for 28%, 26%, 13%, and 8%, or a total of 75%, of the variance in the data.11 After using varimax rotation and solving Equation 12.29, we plotted the first two factor scores for each plant in a scatterplot (Figure 12.8). Like PCA, the factor analysis suggests some discrimination among groups: plants from site HD (black points) have low Factor 1 scores, whereas plants from site TJH (light blue points) have low Factor 2 scores.

Principal Coordinates Analysis

Principal coordinates analysis (PCoA) is a method used to ordinate data using any measure of distance. Principal component analysis and factor analysis are used when we analyze quantitative multivariate data and wish to preserve Euclidean distances between observations. In many cases, however, Euclidean distances between observations make little sense. For example, a binary presence-absence matrix is a very common data structure in ecological and environmental studies: each row of the matrix is a site or sample, and each col11

Some factor analysis software use a goodness-of-fit test (see Chapter 11) to test the hypothesis that the number of factors used in the analysis is sufficient to capture the bulk of the variance versus the alternative that more factors are needed. For the Darlingtonia data, a chi-square goodness-of-fit analysis failed to reject the null hypothesis that four factors were sufficient (χ2 = 17.91, 11 degrees of freedom, P = 0.084).

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High Divide (HD) L.E. Hunter’s Fen (LEH) Day’s Gulch (DG) T.J. Howell’s Fen (TJH)

Factor 2

Figure 12.8 Plot illustrating the first two factors resulting from a factor analysis of the data in Table 12.3. Each point represents an individual plant, and the colors represent the different sites as in Figure 12.6. Results are qualitatively similar to the principal component analysis shown in Figure 12.6, with lower scores for HD on Factor 1 and lower scores for TJH on Factor 2.

–2

–4 –3

–2

–1

0 Factor 1

umn is a species or taxon. The matrix entries indicate the absence (0) or presence (1) of a species in a site. As we saw earlier (see Tables 12.4 and 12.5), Euclidean distances measured for such data may give counterintuitive results. Another example for which Euclidean distances are not appropriate is a genetic distance matrix of the relative differences in electrophoretic bands. In both of these examples, PCoA is more appropriate than PCA. Principal component analysis itself is a special case of PCoA: the eigenvalues and associated eigenvectors of a PCA that we calculated from the variance-covariance matrix also can be recovered by applying PCoA to a Euclidean distance matrix. BASIC CALCULATIONS FOR A PRINCIPAL COORDINATES ANALYSIS

Principal coordi-

nates analysis follows five steps: 1. Generate a distance or dissimilarity matrix from the data. This matrix, D, has elements dij that correspond to the distance between two samples i and j. Any of the distance measurements given in Table 12.6 can be used to calculate the dij’s. D is a square matrix, with the number of rows i = number of columns j = number of observations m. 2. Transform D into a new matrix D*, with elements 1 di*, j = − di2, j 2

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TABLE 12.9 Presence-absence matrix used for principal coordinates analysis (PCoA), correspondence analysis (CA), and non-metric multidimensional scaling (NMDS) Aphaenogaster

CT MA mainland MA islands Vermont

1 1 0 1

Brachymyrmex

Camponotus

Crematogaster

Dolichoderus

Formica

Lasius

0 1 0 0

1 1 0 1

0 0 1 0

0 1 1 0

0 1 1 1

1 1 1 1

Each row represents samples from different locations: Connecticut (CT), the Massachusetts (MA) mainland, islands off the coast of Massachusetts, and Vermont (VT). Each column represents a different ant genus collected during a survey of forest uplands surrounding small bogs. (Data compiled from Gotelli and Ellison 2002a,b and unpublished.)

This transformation converts the distance matrix into a coordinate matrix that preserves the distance relationship between the transformed variables and the original data. 3. Center the matrix D* to create the matrix D with elements δi,j by applying the following transformation to all the elements of D*: δ ij = dij* − di* − d j* + d * – – where d i* is the mean of row i of D*, d j* is the mean of column j of D*, –* and d is the mean of all the elements of D*. 4. Compute the eigenvalues and eigenvectors of D. The eigenvectors ak must be scaled to the square root of their corresponding eigenvalues: a Tk a k = λ k 5. Write the eigenvectors ak as columns, with each row corresponding to an observation. The entries are the new coordinates of the objects in principal coordinates space, analogous to the principal component scores. A BRIEF EXAMPLE We use PCoA to analyze a binary presence-absence matrix of four sites (Connecticut, Vermont, Massachusetts mainland, and Massachusetts Islands) and 16 genera of ants found in forested uplands surrounding small bogs (Table 12.9). We first generated a distance matrix for the four sites using Sørensen’s measure of dissimilarity (Table 12.10). The 4 × 4 distance matrix contains the values of Sørensen’s measure of dissimilarity calculated for all pair-wise combinations of sites. We then computed the eigenvectors, which are used to

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Leptothorax

Myrmecina

Myrmica

Nylanderia

Prenolepis

Ponera

Stenamma

1 1 1 1

0 1 1 0

1 1 1 1

0 0 1 0

0 1 1 0

0 0 0 1

1 1 1 1

Stigmatomma Tapinoma

0 1 1 0

compute the principal coordinate scores for each site. Principal coordinate scores for each site are plotted for the first two principal coordinate axes (Figure 12.9). Sites differ based on dissimilarities in their species compositions, and can be ordinated on this basis. The sites are ordered CT, VT, MA mainland, MA islands from left to right along the first principal axis, which accounts for 80.3% of the variance in the distance matrix. The second principal axis mostly separates the mainland MA sites from the island MA sites and accounts for an additional 14.5% of the variance in the distance matrix. Correspondence Analysis

Correspondence analysis (CA), also known as reciprocal averaging (RA) (Hill 1973b) or indirect gradient analysis, is used to examine the relationship of species assemblages to site charcteristics. The sites usually are selected to span an environmental gradient, and the underlying hypothesis or model is that species abundance distributions are unimodal and approximately normal (or Gaussian) across the environmental gradient (Whittaker 1956). The axes resulting from a TABLE 12.10 Measures of dissimilarity used in principal coordinates analysis (PCoA) and non-metric multidimensional scaling (NMDS)

CT MA mainland MA islands VT

MA mainland

MA islands

0 0.37 0.47 0.13

0.37 0 0.25 0.33

0.47 0.25 0 0.43

0.13 0.33 0.43 0

The original data consist of a presence-absence matrix of forest ant genera in four New England sites (see Table 12.9). Each entry is the Sørensen’s dissimilarity between each pair of sites (see Table 12.6 for formula). The more two sites differ in the genera they contain, the higher the dissimilarity. By definition, the dissimilarity matrix is symmetric and the diagonal elements equal 0.

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Principal coordinate axis 2

422

MA mainland 40

20 MA islands

CT VT

0 0

40 80 Principal coordinate axis 1

Figure 12.9 Ordination of four upland sites based on their ant species assemblages (see Table 12.9) using Principal Coordinates Analysis (PCoA). A pairwise distance matrix for the four sites is calculated from the data in Table 12.10 and used in the PCoA. The sites are ordered CT, VT, MA mainland, and MA islands from left to right along the first principal axis, which accounts for 80.3% of the variance in the distance matrix. The second principal axis mostly separates the mainland MA sites from the island MA sites, and accounts for an additional 14.5% of the variance.

CA maximize the separation of species abundances along each axis of peaks. Correspondence analysis takes as input a site × species matrix that is manipulated as if it were a contingency table (see Chapter 11). Correspondence analysis also can be considered a special case of PCoA; a PCoA using a chi-square distance matrix results in a CA. Correspondence analysis is not much more complex to interpret than PCA or factor analysis, but it requires more matrix manipulations. BASIC CALCULATIONS FOR A CORRESPONDENCE ANALYSIS

1. Begin with a row × column table (such as a contingency table or a site [rows] × species [columns] matrix) with m rows and n columns, and for which m ≥ n. This is often a restriction of CA software. Note that if m ≤ n in the original data matrix, transposing it will meet the condition that m ≥ n, and the interpretation of the results will be identical. 2. Create a matrix Q whose elements qij are proportional to chi-square values:

q i, j =

⎛ observed − expected ⎞ ⎜ ⎟ expected ⎝ ⎠ grand total

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4. 5.

The expected values are calculated as we described in Chapter 11 for chi-square tests for independence (see Equation 11.4). Apply singular-value decomposition to Q: Q = UWVT, where U is an m × n matrix, V is an n × n matrix, U and V are orthonormal matrices, and W is a diagonal matrix (see Appendix for details). Observe that the product QTQ = VWTWVT. The new diagonal matrix WTW, written as L, has elements λi that are the eigenvalues of QTQ. The matrix V has columns that are the eigenvectors in which elements are the loadings for the columns of the original data matrix. The matrix U has columns that are the eigenvectors for which the elements are the loadings for the rows of the original data matrix. Use matrices U and V to separately plot the locations of the rows and columns in ordination space. The locations also can be plotted together in an ordination bi-plot so that you can see relationships between, say, sites and species. However, redundancy analysis (RDA; see below) is a more direct method for analyzing joint relationships between site (environmental) and species (compositional) data.

Like PCA and factor analysis, CA results in principal axes and scores, although in this case we get scores for both the rows (e.g., sites) and columns (e.g., species). As with PCA and factor analysis, the first axis from a CA has the largest eigenvalue (explains the most variance in the data); it also maximizes the association between rows and columns. Subsequent axes account for the residual variation and have successively smaller eigenvalues. Rarely do we use more than two or three axes from CA, because each axis is thought to represent a particular environmental gradient, and a system with more than three unrelated gradients is difficult to interpret. We use CA to examine the joint relationship of ant genera composition at the four sites in Connecticut, Massachusetts, and Vermont (see Table 12.9). These sites differ slightly in latitude (≈3°) and whether they are on the mainland or on islands. The first two axes of the CA account for 58% and 28% (total = 86%) of the variance in the data, respectively. The graph of the ordination of sites (Figure 12.10A) shows discrimination among the sites similar to that observed with the PCoA: CT and VT group together, and are separated from MA along the first principal axis. The second principal axis separates the MA mainland from the MA island sites. Separation of genera with respect to sites is also apparent (Figure 12.10B). Unique genera (such as Ponera in the VT sample, Brachymyrmex in the MA mainland sample, and Crematogaster and Paratrechina in the MA A BRIEF EXAMPLE

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(A)

(B) 100

100

Brachymyrmex

CA axis 2

MA mainland 80

Leptothorax Myrmecina Prenolepis Stigmatomma

Formica

Dolichoderus Lasius Aphaenogaster Myrmica Stenamma Camponotus Ponera

MA islands 0

Crematogaster Nylanderia

Tapinoma

0 20

60 CA axis 1

100

Figure 12.10 Results of a correspondence analysis (CA)

on the site × ant genus matrix in Table 12.9. Plot A shows the ordination of sites, plot B the ordination of genera, and plot C is a bi-plot showing the relationship between the two ordinations. The results of the site ordination are similar to those of the principal coordinates analysis (see Figure 12.9), whereas the ordination of genera separates unique genera such as Ponera in the VT sample, Brachymyrmex in the MA mainland sample, and Crematogaster and Paratrechina in the MA island sample. The ordination results have been scaled so that both the site and species ordinations can be shown on the same plot. In (C), sites are indicated by black dots and ant genera by blue dots.

100

40 60 CA axis 1

100

60 40 20 0

island sample) separate out in the same way as the sites. The remaining genera are arrayed more toward the center of the ordination space (Figure 12.10B). The overlay of the two ordinations captures the relationships of these genera and the sites (compare the site × genus matrix in Table 12.9 with Figure 12.10C). Using more information about the sites (Gotelli and Ellison 2002a), we could attempt to infer additional information about the relationships between individual genera and site characteristics. You should be cautious about drawing such inferences, however. The difficulty is that correspondence analysis carries out a simultaneous ordination of the rows and columns of the data matrix, and asks how well the two ordinations are associated with each other. Thus, we expect a fair degree of overlap between the results of the two ordinations (as shown in Figure 12.10C). Because data

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used for CA essentially are the same as those used for contingency table analysis, hypothesis testing and predictive modeling is better done using the methods described in Chapter 11. Correspondence analysis has been used extensively to explore relationships among species along environmental gradients. However, like other ordination methods, it has the unfortunate mathematical property of compressing the ends of an environmental gradient and accentuating the middle. This can result in ordination plots that are curved into an arch or a horseshoe shape (Legendre and Legendre 1998; Podani and Miklós 2002), even when the samples are evenly spaced along the environmental gradient. An example of this is seen in Figure 12.10A. The first axis would array the four sites linearly, and the second axis pulls the MA mainland site up. These four points form a relatively smooth arch. In many instances, the second CA axis simply may be a quadratic distortion of the first axis (Hill 1973b; Gauch 1982). A reliable and interpretable ordination technique should preserve the distance relationships between points; their original values and the values created by the ordination should be equally distant (to the extent possible given the number of axes that are selected) when measured with the same distance measure. For many common distance measures (including the chi-square measure used in CA), the horseshoe effect distorts the distance relationships among the new variables created by the ordination. Detrended correspondence analysis (DCA; Gauch 1982) is used to remove the horseshoe effect of correspondence analysis and presumably illustrate more accurately the relationship of interest. However, Podani and Miklós (2002) have shown that the horseshoe is a mathematical consequence of applying most distance measures to species that have unimodal responses to underlying environmental gradients. Although DCA is implemented widely in ordination software, it is no longer recommended (Jackson and Somers 1991). The use of alternative distance measures (Podani and Miklós 2002) is a better solution to the horseshoe problem.

THE HORSESHOE EFFECT AND DETRENDING

Non-Metric Multidimensional Scaling

The four previous ordination methods are similar in that the distances between observations in multivariate space are preserved to the extent possible after the multivariate data have been reduced to a smaller number of composite variables. In contrast, the goal of non-metric multidimensional scaling (NMDS) is to end up with a plot in which different objects are placed far apart in the ordination space, while similar objects are placed close together. Only the rank ordering of the original distances or dissimilarities is preserved.

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BASIC COMPUTATIONS OF A NON-METRIC MULTIDIMENSIONAL SCALING

Carrying out a non-metric multidimensional scaling requires nine steps, some of which are repeated. 1. Generate a distance or dissimilarity matrix D from the data. Any distance or dissimilarity measure can be used (see Table 12.6). The elements of dij , D are distances or dissimilarities between observations. 2. Choose the number of dimensions (axes) n to be used to draw the ordination. Use two or three axes because most graphs are two- (x- and y-axes) or three- (x-, y-, and z-axes) dimensional. 3. Start the ordination by placing the m observations in the n-dimensional space. Subsequent analyses depend strongly on this initialization, because NMDS finds its solution by local minimization (similar to nonlinear regression; see Chapter 9). If some geographic information is available (such as latitude and longitude), that may be used as a good starting point. Alternatively, the output from another ordination (such as PCoA) can be used to determine the initial positions of observations in an NMDS. 4. Compute new distances δij between the observations in the initial configuration. Normally, Euclidean distances are used to calculate δij. 5. Regress δij on dij. The result of this regression is a set of predicted values δˆij. For example, if we use a linear regression, δij = β0 + β1dij + εij , then the predicted values are δˆ ij = βˆ 0 + βˆ 1dij 6. Compute a goodness of fit between δij and δˆij. Most NMDS programs compute this goodness of fit as a stress: m

Stress =

∑ ∑ (δ ij − δˆ ij )2 i=1 j =1 m

∑ ∑ δˆ ij2 i=1 j =1

Stress is computed on the lower triangle of the square D matrix. The numerator is the grand sum of the square of the difference between observed and expected values, and the denominator is the grand sum of the squared expected values. Note that the stress looks a lot like a chisquare test-statistic (see Chapter 11).

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7. Change the position of the m observations in n-dimensional space slightly in order to reduce the stress. 8. Repeat steps 4–7 until the stress can no longer be reduced any further. 9. Plot the position of the m observations in n-dimensional space for which stress is minimal. This plot illustrates “relatedness” among observations. We use NMDS to analyze the ant data in Table 12.8. We used Sørensen’s measure of dissimilarity for our distance measurement. The scree plot of stress scores indicates that two dimensions are sufficient for the analysis (Figure 12.11). As with the CA and PCoA, we have good discrimination among the sites (Figure 12.12A), and good separation among unique assemblages of genera (Figure 12.12B). Because scaling of axes in NMDS is arbitrary, we cannot overlay these two plots in an ordination bi-plot.

A BRIEF EXAMPLE

Advantages and Disadvantages of Ordination

Ecologists and environmental scientists use ordination to reduce complex multivariate data to a smaller, more manageable set of data; to sort sites on the basis of environmental variables of species assemblages; and to identify species responses to environmental gradients or perturbations. Because some ordination methods (such as PCA and factor analysis) are commonly available in most commercial statistical packages, we have easy, often menu-driven access to these tools. Used prudently, ordination can be a powerful tool for exploring data, illustrating patterns, and generating hypotheses that can be tested with subsequent sampling or experiments. The disadvantages of ordination are not apparent because these techniques have been automated in many software packages, the mechanics are hidden, and the assumptions are rarely spelled out. Ordination is based on matrix algebra, 40

Stress

Figure 12.11 Scree plot for dimensions in a

0 1

3 Dimensions

NMDS analysis of the site × ant genus data (see Table 12.9). A pairwise distance matrix for the four sites is calculated from these data (see Table 12.10) and used in the NMDS. In an NMDS, the stress in the data is a deviation measure similar to a chi-square goodness-of-fit statistic. This scree plot illustrates the successive reduction in stress with increasing dimension in the NMDS of the ant data. No further significant reduction in stress occurred after three dimensions, which account for most of the goodness-of-fit.

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(A)

(B) Ponera Dolichoderus Lasius Myrmica Stenamma

NMDS axis 2

CT MA islands

MA mainland NMDS axis 1

Tapinoma Crematogaster Nylanderia

Aphaenogaster Camponotus

Formica Leptothorax Myrmecina Prenolepis Stigmatomma

Brachymyrmex

NMDS axis 1

Figure 12.12 Plot of the first two dimensions of the NMDS analysis of the site × ant genus data in Table 12.9. A pairwise distance matrix for the four sites is calculated from the data in Table 12.10 and is used in the NMDS. (A) Ordination of sites. (B) Ordination of genera.

with which many ecologists and environmental scientists are unfamiliar. Because principal axes normally are rescaled prior to plotting principal scores, the scores are interpretable only relative to each other and it is difficult to relate them back to the original measurements. RECOMMENDATIONS It is rarely obvious which ordination technique you should choose to best order observations, samples, or populations in multivariate space. Simple data reduction is best accomplished by principal component analysis (PCA) when Euclidean distances are appropriate and outliers or highly skewed data are not present. Principal coordinates analysis (PCoA) is appropriate when other distance measures are needed; PCA is equivalent to a PCoA when Euclidean distances are used. We recommend PCoA for most ordination applications in which the goal is to preserve the original multivariate distances between observations in the reduced (ordination) space. Non-metric multidimensional scaling (NMDS) only preserves the rank ordering of the distances, but it can be used with any distance measure. Correspondence analysis (CA) is a reasonable method of ordination for examining species distributions along environmental gradients, but its cousin, detrended correspondence analysis (DCA), should no longer be used. Likewise, the results of factor analysis are difficult to interpret and too dependent on subjective rotation methods. In all cases, it is important to remember the underlying ecological or environmental question of interest. For example, if the original question was to see how species or characteristics are dis-

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tributed along an environmental gradient, then simply plotting the response variable against the appropriate measurement of the environment (or the first or second principal axes) may be more informative than any ordination plot. Whichever method is employed, ordination should be used cautiously for testing hypotheses; it is best used for data exploration and pattern generation. However, classical hypothesis testing can be carried out on scores resulting from any ordination, as long as the assumptions of the tests are met.

Classification Classification is the process by which we place objects into groups. Whereas the goal of ordination is to separate observations or samples along environmental gradients or biological axes, the goal of classification is to group similar objects into identifiable and interpretable classes that can be distinguished from neighboring classes. Taxonomy is a familiar application of classification—a taxonomist begins with a collection of specimens and must decide how to assign them to species, genera, families, and higher taxonomic groups. Regardless of whether taxonomy is based on morphology, gene sequences, or evolutionary history, the goal is the same: an inclusive classification system based on hierarchical clusters of groups. Ecologists and environmental scientists may be suspicious of classifications because the classes are assumed to represent discrete entities, whereas we are more used to dealing with continuous variation in community structure or morphology that maps onto continuous environmental gradients. Ordination identifies patterns occurring within gradients, whereas classification identifies the endpoints or extremes of the gradients while ignoring the in-between. Cluster Analysis

The most familiar type of classification analysis is cluster analysis. Cluster analysis takes m observations, each of which has associated with it n continuous numerical variables, and segregates the observations into groups. Table 12.11 illustrates the kind of data used in cluster analysis. Each of the nine rows of the table represents a different country in which samples were collected. Each of the three columns of the table represents a different morphological measurement taken on specimens of the marine snail Littoraria angulifera. The goal of this analysis is to form clusters of sites on the basis of similarity in shell shape. Cluster analysis also can be used to group sites on the basis of species abundances or presences-absences, or to group organisms on the basis of similarity in measured characteristics such as morphology or DNA sequences.

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TABLE 12.11 Snail shell measurements used for cluster analysis Country

Continent

Proportionality

Circularity

Spire Height

Angola Bahamas Belize Brazil Florida Haiti Liberia Nicaragua Sierra Leone

Africa N. America N. America S. America N. America N. America Africa N. America Africa

1.36 1.51 1.42 1.43 1.45 1.49 1.36 1.48 1.35

0.76 0.76 0.76 0.74 0.74 0.76 0.75 0.74 0.73

1.69 1.86 1.85 1.71 1.86 1.89 1.69 1.69 1.72

Shell shape in the snail Littoraria angulifera was measured for samples from 9 countries. Shell proportionality = shell height/shell width; shell circularity = aperture width/aperture height; and spire height = shell height/aperture length. Values in the table are averages based on 2 to 100 samples per site (data from Merkt and Ellison 1998). Cluster analysis groups the different countries based on similarity in shell morphology (see Figure 12.13).

Choosing a Clustering Method

There are several methods available for clustering data (Sneath and Sokal 1973), but we will focus on only two that are commonly used by ecologists and environmental scientists. Agglomerative clustering proceeds by taking many separate observations and grouping them into successively larger clusters until one cluster is obtained. Divisive clustering, on the other hand, proceeds by placing all the observations in one group and then splitting them into successively smaller clusters until each observation is in its own cluster. For both methods, a decision must be made, often based on some statistical rule, as to how many clusters to use in describing the data. We illustrate these two methods using the data in Table 12.11. Agglomerative methods begin with a square m × m distance matrix, in which the entries are the pairwise morphological distances measured for each pair of sites; Euclidean distances are the most straightforward to use (Table 12.12), but any distance measure (see Table 12.6) can be used in cluster analysis. An agglomerative cluster analysis starts with all the objects—sites in this analysis—separately and successively groups them. We see from the Euclidean distance matrix (see Table 12.12) that Brazil is nearest (in distance units) to Nicaragua, Angola is nearest to Liberia, Belize is nearest to Florida, and The Bahamas are nearest to Haiti. AGGLOMERATIVE VERSUS DIVISIVE CLUSTERING

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TABLE 12.12 Euclidean distance matrix for morphological measurements of snail shells Angola

Angola Bahamas Belize Brazil Florida Haiti Liberia Nicaragua Sierra Leone

0 0.23 0.17 0.08 0.19 0.24 0.01 0.12 0.04

Bahamas Belize Brazil

0 0.09 0.17 0.06 0.04 0.23 0.17 0.21

0 0.14 0.04 0.08 0.17 0.17 0.15

0 0.15 0.19 0.07 0.05 0.08

Florida Haiti

0 0.05 0.19 0.17 0.17

0 0.24 0.20 0.22

Liberia

0 0.12 0.04

Nicaragua Sierra Leone

0 0.13

The original data are detailed in Table 12.11. Because distance matrices are symmetrical, only the lower half of the matrix is shown.

These clusters form the bottom row of the dendrogram (tree-diagram) shown in Figure 12.13. In this example, Sierra Leone is the odd site out (agglomerative clustering works upwards in pairs), and its addition to the Angola-Liberia cluster to form a new African cluster. The resulting four new clusters are clustered further in such a way that successively larger clusters contain smaller ones. Thus, the Bahamas–Haiti cluster is joined with the Belize–Florida cluster to form a new Caribbean–North Atlantic cluster; the other two clusters form a South Atlantic–Africa cluster. A divisive cluster analysis begins with all the objects in one group, and then divides them up based on dissimilarity. In this case, the resulting clusters are the same ones that were produced with agglomerative clustering. More typically, divisive clustering results in fewer clusters, each with more objects, whereas agglomerative clustering results in more clusters, each with fewer objects. Divisive clustering algorithms also may proceed to a predetermined number of clusters that is specified by the investigator in advance of performing the analysis. A now rarely used method for community classification, TWINSPAN (for “two-way indicator-species analysis”; see Gauch 1982), uses divisive clustering algorithms. HIERARCHICAL VERSUS NON-HIERARCHICAL METHODS Intuitively, clusters with few observations should be embedded within higher-order clusters with more observations. In other words, if observations a and b are in one cluster, and observations c and d are in another cluster, a larger cluster that includes a and c should also include b and d. This classification arrangement should be familiar from taxonomy: if two species are in one genus, and two other species are in anoth-

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0.20

0.15 Euclidean distance

0.10

Nicaragua

Brazil Liberia

Angola

Sierra Leone

Florida

Belize

0.0

Haiti

0.05

Bahamas

432

Figure 12.13 Results of an agglomerative cluster analysis. The original data (Merkt and Ellison 1998) consist of morphological ratios of snail (Littoraria angulifera) shell measurements from several countries (see Table 12.11). A dendrogram (branching figure) groups the sites into clusters based on similarity in shell morphology. The y-axis indicates the Euclidean distance for which sites (or lower-level clusters) are joined into a higher-level cluster.

er genus, and both genera are in the same family, then all four species are also in that same family. Clustering methods that follow this rule are called hierarchical clustering methods. Both agglomerative and divisive clustering methods can be hierarchical. In contrast, non-hierarchical clustering methods group observations independently of an externally imposed reference system. Ordination can be considered analogous to non-hierarchical clustering—the results of an ordination often are independent of externally imposed orderings of the system. For example, two species in the same genus may wind up in different clusters produced by ordination of a set of morphological or habitat characteristics. Ecologists and environmental scientists generally use non-hierarchical methods to search for patterns in the data and to generate hypotheses that should then be tested with additional observations or experiments.

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K-means clustering is a non-hierarchical method that requires you to first specify how many clusters you want. The algorithm creates clusters in such a way that all the objects within one cluster are closer to each other (based on a distance measure) than they are to objects within the other clusters. K-means clustering minimizes the sums of squares of distances from each object to the centroid of its cluster. We first applied K-means clustering to the snail data by specifying two clusters. One cluster, consisting of samples from Angola, Brazil, Liberia, Nicaragua, and Sierra Leone, had relatively small shells (centroid: proportionality = 1.39, circularity = 0.74, spire height = 1.70), and the other cluster (Bahamas, Belize, Florida, and Haiti) had relatively large shells (centroid = 1.47, 0.76, 1.87). The within-cluster sum of squares for the first (African–South Atlantic) cluster equals 0.14, and the within-cluster sum of squares for the second (Caribbean–North Atlantic) cluster equals 0.06. A K-means clustering of these data specifying three clusters split the African–South Atlantic cluster into two clusters, one for the African sites and one for Nicaragua and Brazil. How do you choose how many clusters to use? There is no hard-and-fast rule, but the more clusters used, the fewer members in each cluster, and the more similar the clusters will be to one another. Hartigan (1975) suggests you first perform a K-means clustering with k and k + 1 clusters on a sample of m observations. If ⎡ ∑ SS(within k clusters ) ⎤ × (m − k − 1)⎥ >10 ⎢ ⎣ ∑ SS( within k+1 clusters ) ⎦

(12.30)

then it makes sense to add the (k + 1)th cluster. If the inequality in Equation 12.30 is less than 10, it is better to stop with k clusters. Equation 12.30 will generate a large number whenever adding a cluster substantially increases the within-group sum of squares. However, in simulation studies, Equation 12.30 tends to overestimate the true number of clusters (Sugar and James 2003). For the snail data in Table 12.11, Equation 12.30 = 8.1, suggesting that two clusters is better than three. Sugar and James (2003) review a wide range of methods for identifying the number of clusters in a dataset, an active area of research in multivariate analysis. Discriminant Analysis

Discriminant analysis is used to assign samples to groups or classes that are defined in advance; it behaves like a cluster analysis in reverse. Continuing with our snail example, we can use the geographic locations to define groups

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in advance. The snail samples come from three continents: Africa (Angola, Liberia, and Sierra Leone), South America (Brazil), and North America (all the rest). If we were given only the shell shape data (three variables), could we assign the observations to the correct continent? Discriminant analysis is used to generate linear combinations of the variables (as in PCA; see Equation 12.21) that are then used to separate the groups as best as possible. As with MANOVA, discriminant analysis requires that the data conform to a multivariate normal distribution. We can generate a linear combination of the original variables using Equation 12.21 Zi = ai1Y1 + ai2Y2 + … + ainYn — If the multivariate means Y vary among groups, we want to find the coefficients a in Equation 12.20 that maximize the differences among groups. Specifically, the coefficients ain are those that would maximize the F-ratio in an ANOVA— in other words, we want the among-group sums of squares to be as large as possible for a given set of Zi’s. Once again, we turn to our sums of squares and cross-products (SSCP) matrices H (among-groups) and E (within-groups, or error; see the discussion of MANOVA earlier in the chapter). The eigenvalues and eigenvectors are then determined for the matrix E –1H. If we order the eigenvalues from largest to smallest (as in PCA, for example), their corresponding eigenvectors ai are the coefficients for Equation 12.21. The results of this discriminant analysis are given in Table 12.13. This small subset of the full dataset departs from multivariate normality (En = 31.32, P = 2 × 10–5, with 6 degrees of freedom), but the full dataset (of 1042 observations) passed this test of multivariate normality (P = 0.13). Although the multivariate means (Table 12.13A) do not differ significantly by continent (Wilk’s lambda = 0.108, F = 2.7115, P = 0.09 with 6 and 8 degrees of freedom), we can still illustrate the discriminant analysis.12 The first two eigenvectors (Table 12.13B) accounted for 97% of the variance in the data, and we used these eigenvectors in Equation 12.21 to calculate principal component scores for each of the original samples. These scores for the nine observations in Table 12.11 are plotted in Figure 12.14, with different colors corresponding to shells from different con-

12 The full dataset analyzed by Merkt and Ellison (1998) had 1042 samples from 19 countries. Discriminant analysis was used to predict from which oceanographic current system (Caribbean, Gulf of Mexico, Gulf Stream, North or South Equatorial) a shell came from. The discriminant analysis successfully assigned more than 80% of the samples to the correct current system.

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TABLE 12.13 Discriminant analysis by continent of snail shell morphology data from 9 sites A. Variable means

Africa N. America S. America Proportionality 1.357 1.470 1.430 Circularity 0.747 0.752 0.740 Spire height 1.700 1.830 1.710 The first step is to calculate the variable means for the samples grouped by each continent. B. Eigenvectors

a1 a2 Proportionality 0.921 0.382 Circularity –0.257 –0.529 Spire height 0.585 –0.620 Discriminant analysis generates linear combinations of these variables (similar to PCA) that maximally separate the samples by continent. The eigenvectors contain the coefficients (loadings) that are used to create a discriminant score for each sample; only the first two, which explain 97% of the variance, are shown. C. Classification matrix

Predicted (classified) Africa N. America S. America % Correct Observed Africa 3 0 0 100 N. America 0 4 1 80 S. America 0 0 1 100 The samples are assigned to one of the three continents according to their discriminant scores. In this classification matrix, all but one of the samples was correctly assigned to its original continent. This result is not too surprising because the same data were used to create the discriminant score and to classify the samples. D. Jackknifed classification matrix

Predicted (classified) Africa N. America S. America % Correct Observed Africa 2 0 1 67 N. America 0 3 2 60 S. America 0 1 0 0 A more unbiased approach is to use a jackknifed classification matrix in which the discriminant score is created from m – 1 samples and then used to classify the excluded sample. In this way, the discriminant function is independent of the sample being classified. The jackknifed classification did not perform nearly as well as the non-jacknifed matrix, with only 5 of the 9 samples correctly classified by continent. The full data from which these ratios are drawn is detailed in Table 12.11. Discriminant analyses and other multivariate techniques should be based on much larger samples than we have used in this example. (Data from Merkt and Ellison 1998.)

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tinents. We see an obvious cluster of the African samples, and another obvious cluster of the North American samples. However, one of the North American samples is closer to the South American sample than it is to the other North American samples. The predicted classification is based on minimizing distances (usually the Mahalanobis or Euclidean distance; see Table 12.6) among the observations within groups. Applying this method to the data results in accurate predictions for the South American and African shells, but only 80% (4 of 5) of the North American shells are correctly classified. Table 12.13C summarizes these classifications as a classification matrix. The rows of the square classification matrix represent the observed groups and the columns represent the predicted (classified) groups. The entries represent the number of observations that were classified into a particular group. If the classification algorithm has no errors, then all of the observations will fall on the diagonal. Off-diagonal values represent errors in classification in which an observation was assigned incorrectly. It should come as no surprise that the predicted classification in a discriminant analysis closely matches the observed data; after all, the observed data were used to generate the eigenvectors that we then used to classify the data! Thus, we expect the results to be biased in favor of correctly assigning our observations to the groups from which they came. One solution to overcoming this bias is to jackknife the classification matrix in Table 12.13C. Jack-

South America North America Africa

Figure 12.14 A plot of the first two axes of a discriminant analysis on morphological ratios of snail (Littoraria angulifera) shell measurements from several countries (see Table 12.11). Discriminant analysis is used to evaluate how well the shells from the three different continents can be distinguished. Discriminant scores were calculated using the eigenvectors in Table 12.13 and Equation 12.21.

Discriminant score 2

–0.88

–0.92

–0.96

–1.00

2.00

2.05

2.10

2.15 2.20 Discriminant score 1

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2.30

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knifing proceeds by dropping one individual observation from the dataset, re-analyzing the data without that observation, and then using the results to classify the dropped observation (see also “The Influence Function” in Chapter 9, and Footnote 2 in Chapter 5). The jackknifed classification matrix is given in Table 12.13D. This classification is much poorer, because the multivariate means for each continent were not very different. On the other hand, we normally would not conduct a discriminant analysis on such a small dataset. Another solution, if the original dataset is large, is to partition it randomly into two groups. Use one group to construct the classification and use the other to test it. New data can also be collected and classified with the discriminant function, although there needs to be some way to independently verify that the group assignments are correct. Advantages and Disadvantages of Classification

Classification techniques are relatively easy to use and interpret, and are implemented widely in statistical software. They allow us either to group samples, or to assign samples to groups identified a priori. Dendrograms and classification matrices clearly summarize and communicate the results of cluster analysis and discriminant analysis. Both cluster analysis and discriminant analysis must be used cautiously, however. There are many ways to carry out a cluster analysis, and different methods usually yield different results. It is most important to decide on a method beforehand instead of trying all the different methods and picking the one that looks the nicest or conforms to preconceived notions. Discriminant analysis is much more straightforward to perform, but it requires both ample data and assignment of groups a priori. Both cluster analysis and discriminant analysis are descriptive and exploratory methods. The MANOVA statistics described earlier in this chapter can be used on the results of a discriminant analysis to test hypotheses about differences among groups. Conversely, discriminent analysis can also be used as an a posteriori test to compare groups once the null hypothesis in a MANOVA has been rejected. Indeed, cluster analysis and discriminant analysis begin with the assumption that distinct groups or clusters exist, whereas a proper null hypothesis would be that the samples are drawn from a single group and exhibit only random differences among one another. Strauss (1982) describes Monte Carlo methods for testing the statistical significance of clusters. Bootstrapping and other computer-intensive methods for testing the statistical significance of clusters are also widely used in phylogenetic analysis (Felsenstein 1985; Emerson et al. 2001;

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Huelsenbeck et al. 2002; Sanderson and Shaffer 2002; Miller 2003; Sanderson and Driskell 2003).

Multivariate Multiple Regression The methods for multivariate analysis described thus far have been primarily descriptive (ordination, classification) or limited to categorical predictor variables (Hotelling’s T 2 test and MANOVA). The final topic for this chapter is the extension of regression (one continuous response variable and one or more continuous predictor variables) to multivariate response data. Two methods are used commonly by ecologists: redundancy analysis (RDA) and canonical correspondence analysis (CCA). We only describe the mechanics of RDA, which is the direct extension of multiple regression to the multivariate case. RDA assumes a causal relationship between the independent and dependent variables, whereas CCA focuses on generating a unimodal axis with respect to the response variables (e.g., species occurrences or abundances) and a linear axis with respect to the predictor variables (e.g., habitat or environmental characteristics). A third technique, canonical correlation analysis (Hotelling 1936; Manly 1991) is based on symmetrical relationships between the two variables. In other words, canonical correlation analysis assumes error in both the predictor and response (see discussion of the assumptions of regression in Chapter 9). Redundancy Analysis

One of the most common uses of RDA is to examine relationships between species composition, measured as the abundances of each of n species, and environmental characteristics, measured as a set of m environmental variables (Legendre and Legendre 1998). The species composition data represent the multivariate response variable, and the environmental variables represent the multivariate predictor variable. But RDA is not restricted to that kind of analysis— it can be used for any multiple regression involving multiple response and multiple predictor variables, as we demonstrate in our example. THE BASICS OF A REDUNDANCY ANALYSIS

As developed in Chapter 9, the stan-

dard multiple linear regression is Y j = β 0 + β1 X1 + β 2 X 2 + … + β m X m + ε j

(12.31)

The βi values are the parameters to be estimated, and εj represents the random error. Standard least squares methods are used to fit the model and provide unbiased estimates bˆ and eˆ of the βi’s and εj’s . In the multivariate case, the sin-

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Multivariate Multiple Regression

gle response variable Y is replaced by a matrix of n variables Y. Redundancy analysis regresses Y on a matrix of independent variables X, all measured for each observation. The steps of an RDA are: 1. Regress each of the individual response variables Yj in Y on all the variables in X (using Equation 12.31). This results in a matrix of fitted values Yˆ = ⎡⎣Yˆj ⎤⎦ . This matrix is calculated as Yˆ = XB, where B is the matrix of regression coefficients: B = (X T X )−1 X T Y 2. Use the Yˆ matrix in a principal component analysis of its standardized sample variance-covariance matrix, yielding a matrix A of eigenvectors. The eigenvectors have the same interpretation as they do in a PCA. If you are performing an RDA on a matrix of environmental characteristics × species, the elements of A are called the species scores. 3. Generate two sets of scores from this PCA. a. The first set of scores, F, are computed by multiplying the matrix of eigenvectors A by the matrix of response variables Y. The result, F = YA, is a matrix in which the columns are called site scores. The site scores F are an ordination relative to Y. If you are using an environmental characteristics × species matrix, the site scores are based on the original, observed species distributions. b. The second set of scores, Z, are computed by multiplying the matrix of eigenvectors A by the matrix of fitted values Yˆ. The result, Z = Yˆ A, is a matrix in which the columns are called fitted site scores. Because Yˆ is also equal to the matrix XB, the matrix of fitted site scores can also be written as Z = XBA. The fitted site scores Z are an ordination relative to X. If you are using an environmental characteristics × species matrix, the fitted site scores are based on the distribution of species that are predicted by the environment. 4. Next we want to know how the predictor variables in X contribute to each of the ordination axes. The most straightforward way to measure the contribution of the predictor variables is to observe that the Z matrix is composed of two parts: X, the predictor values, and BA, the matrix product of the regression coefficients and the eigenvectors of the Yˆ matrix. The matrix C = BA tells us the contribution of the X variables to the matrix of fitted site scores Z. This decomposition is equivalent to saying that the values in each of the columns of C are equal to standardized regression coefficients on the matrix X. An alternative method is to determine the correlation between the environmental (predictor) vari-

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ables X and the site scores F, which is the product of the response variables and their eigenvectors. 5. Finally, we construct a bi-plot, as we did for corresponding analysis, in which we plot the principal axes of either F (the observed) or Z (the fitted) scores for the predictor variables. On this plot, we can place the locations of the response variables. In this way, we can visualize the multivariate relationship of the response variables to the predictor variables. To illustrate RDA, we return to the snail data (see Table 12.11). For each of these sites we also have measures of four environmental variables: annual rainfall (mm), number of dry months per year, mean monthly temperature, and mean height of the mangrove canopy in which the snails foraged (Table 12.14). The results of the four steps of the RDA are presented in Table 12.15. Because the response variables are measures of shell-shape, the A matrix is composed of shell-shape scores. The F and Z matrices are site scores and fitted site scores, respectively. Figure 12.15 illustrates how sites co-vary with environmental variables (blue points and directional arrows) and how shell shapes co-vary with the sites and their environmental variables (black points and directional arrows). For example, the Brazilian, Nicaraguan, and African sites are characterized by greater canopy height and greater rainfall, whereas the Florida and Caribbean sites are characterized by higher temperatures and longer dry months. Shells from the Caribbean and Florida have larger proportionality and higher spire heights, whereas shells from Africa, Brazil, and Nicaragua are more circular. A BRIEF EXAMPLE

TABLE 12.14 Environmental variables used in the redundancy analysis (RDA) Country

Angola Bahamas Belize Brazil Florida Haiti Liberia Nicaragua Sierra Leone

Annual rainfall (mm)

No. dry months

Mean monthly temp. (oC)

Mean height of forest canopy (m)

363 1181 1500 2150 1004 1242 3874 3293 4349

9 2 2 4 1 6 3 0 4

26.4 25.1 29.5 26.4 25.3 27.5 27.0 26.0 26.6

30 3 8 30 10 10 30 15 35

Data from Merkt and Ellison (1998).

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TABLE 12.15 Redundancy analysis of small shell morphology, and environmental data A. Compute the matrix of fitted values Yˆ = X(X T X )−1 X T Y.

⎡−0.05 ⎢ ⎢ 0.09 ⎢ 0.00 ⎢ ⎢−0.05 Yˆ = ⎢ 0.05 ⎢ ⎢ 0.04 ⎢−0.05 ⎢ ⎢ 0.03 ⎢ ⎣−0.06

0.01 −0.04 ⎤ ⎥ 0.00 0.09⎥ 0.01 0.08⎥ ⎥ −0.01 −0.08⎥ 0.00 0.03⎥ ⎥ 0.02 0.09⎥ −0.01 −0.07 ⎥ ⎥ −0.01 0.00⎥ ⎥ −0.01 −0.10⎦

This matrix is the result of regressing each of the three shape variables on the four environmental variables. Each row of Yˆ is one observation (from one site; see Table 12.11), and each column is one of the variables (proportionality, circularity, and spire height, respectively). B. Use Yˆ in a PCA to compute the eigenvectors A.

⎡0.55 −0.81 −0.19⎤ ⎥ ⎢ A = ⎢0.08 0.27 −0.96⎥ ⎢⎣0.83 0.52 0.21⎥⎦ These are the shell-shape scores, and could be used to calculate principal component values (Zi) for shell shape. Columns represent the first three eigenvectors, and rows represent the three morphological variables. Component 1 accounts for 94% of the variance in shell shape. Shell circularity, which is roughly constant across sites (see Table 12.11), has little weight (a21 = 0.08) on Principal Component 1. C. Compute the matrix of site scores F = YA and the matrix of fitted site scores Z = Yˆ A.

⎡2.21 ⎢ ⎢2.44 ⎢2.38 ⎢ ⎢2.26 F = ⎢2.40 ⎢ ⎢2.45 ⎢2.21 ⎢ ⎢2.28 ⎢ ⎣2.23

−0.63⎤ ⎥ −0.62⎥ −0.61⎥ ⎥ −0.62⎥ −0.59⎥ ⎥ −0.61⎥ −0.62⎥ ⎥ −0.63⎥ ⎥ −0.01 −0.59⎦

−0.03 −0.06 0.00 −0.08 −0.02 −0.03 −0.03 −0.13

0.02 −0.01⎤ ⎡−0.06 ⎥ ⎢ ⎢ 0.13 −0.03 0.00⎥ ⎢ 0.07 0.04 0.01⎥ ⎥ ⎢ ⎢−0.09 0.00 0.00⎥ Z = ⎢ 0.05 −0.02 0.00⎥ ⎥ ⎢ 0.00⎥ ⎢ 0.10 0.02 ⎢−0.09 0.00 0.00⎥ ⎥ ⎢ ⎢ 0.01 −0.03 0.00⎥ ⎥ ⎢ ⎣−0.12 0.00 0.00⎦

Matrix F contains the principal component scores for shell-shape values obtained by multiplying the original values (Y) by the eigenvectors in matrix A. Matrix Z contains the principal component scores for shell-shape values obtained by multiplying the fitted values ( Yˆ ), which are derived from the habitat matrix X (see Table 12.14), by the eigenvectors in A.

(continued)

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TABLE 12.15 (continued) D. Compute the matrix C = BA to yield the regression coefficients on X.

⎡ 0.01 ⎢ 0.03 C=⎢ ⎢ 0.00 ⎢ ⎢⎣−0.11

0.00 0.00⎤ ⎥ 0.01 0.00⎥ 0.02 0.00⎥ ⎥ 0.00 0.00⎥⎦

Rows are the four environmental variables, and columns are the first three components. Coefficients of 0.00 are rounded values of small numbers. Alternatively, we can compute the correlations of the environmental variables X with the site scores F. The cell entries are the correlation coefficients between X and the columns of F: Rainfall Dry months Mean temperature Canopy height

Component 1 –0.55 –0.28 0.02 –0.91

Component 2 –0.19 0.34 0.49 0.04

Component 3 0.11 –0.18 0.11 –0.04

In this example, we have regressed snail shell-shape characteristics (see Table 12.11) on environmental data (see Table 12.13). Prior to analysis, the environmental data were standardized using Equation 12.17. (Data from Merkt and Ellison 1998.)

The results of RDA (and CCA) can be tested for statistical significance using Monte Carlo simulation methods (see Chapter 5). The null hypothesis is that there is no relationship between the predictor variables and the response variables. Randomizations of the raw data— interchanging the rows of the Y matrix—are followed by repetition of the RDA. We then compare an F-ratio computed for the original data with the distribution of F-ratios for the randomized data. The F-ratio for this test is:

TESTING SIGNIFICANCE OF THE RESULTS

⎛ n ⎜ ∑ λj ⎜ j =1 ⎜ m ⎝

⎞ ⎟ ⎟ ⎟ ⎠

(12.32)

⎛ RSS ⎞ ⎜ ⎟ ⎝ ( p − m − 1) ⎠

In this equation, ∑ λ j is the sum of the eigenvalues; RSS is the residual sums of squares (computed as the sum of the eigenvalues of a PCA using (Y – Yˆ ) in place of Yˆ (Table 12.15B); m is the number of predictor variables; and p is the number of observations. Although the sample size is too small for a meaning-

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RDA axis 2

–1

Circularity

Belize

Sierra Leone Angola Liberia

Florida Haiti Proportionality Bahamas

Canopy height Brazil Rainfall

–2

–3 –1.5

Temperature

Dry months

Spire height

Nicaragua

–1.0

–0.5

0.0 0.5 RDA axis 1

1.0

1.5

Figure 12.15 Bi-plot of the first two axes of a Redundancy Analysis (RDA) regressing snail shell data (see Table 12.11) on environmental data (see Table 12.14), all of which were measured for nine countries (Merkt and Ellison 1998). The country labels indicate the placement of each site in the ordination space (the F matrix of Table 12.15). The black symbols indicate the placement of the morphological variables in site space. Thus, shells from Belize, Florida, Haiti, and the Bahamas have larger proportionality and higher spire heights than shells in the other sites, whereas African, Brazilian, and Nicaraguan shells are more circular. Black arrows indicate the direction of increase of the morphological variable. Similarly, the blue points and arrows indicate how the sites were ordinated. All values were standardized to enable plotting of all matrices on similar scales. Thus, the lengths of the arrows do not indicate magnitude of effects. However, they do indicate their directionality or loading on the two axes.

ful analysis, we can nevertheless use Equation 12.32 to see if our results are statistically significant. The F-ratio for our original data = 3.27. A histogram of Fratios from 1000 randomizations of our data is shown in Figure 12.16. Comparison of the observed F-ratio with the randomizations gives an exact P-value of 0.095, which is marginally larger than the traditional P = 0.05 cutoff. The narrow conclusion (based on a limited sample size) is that there is no significant relationship between snail morphology and environmental variation among these nine sites.13 See Legendre and Legendre (1998) for further details and examples of hypothesis testing in RDA and CCA. 13

In contrast, the analysis of the full dataset of 1042 observations from 19 countries yielded a significant association between snail shell shape and environmental conditions (Merkt and Ellison 1998).

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Figure 12.16 Frequency distribution of F-ratios

500 400 Frequency

resulting from 1000 randomizations and subsequent redundancy analysis (RDA) of the snail data (see Table 12.11), in which the country labels have been randomly re-shuffled among the samples. The F-ratio for the observed data = 3.27. Ninety-five of the randomizations had a larger F-ratio, so the P-value for the RDA in Figure 12.15 = 0.095. This result is marginally nonsignificant using an α-level equal to 0.05, reflecting the limitation of using RDA with such a small dataset.

300 200

Fobserved = 3.27

100 0 0

4 5 6 7 Simulated F-ratios

An extension of RDA, distance-based RDA (or db-RDA) is an alternative method of testing complex multivariate models (Legendre and Anderson 1999; McArdle and Anderson 2001). Unlike a classical MANOVA, db-RDA does not require that data be multivariate normally distributed, that distance measurements between observations be Euclidean, or that there be more observations than there are measured response variables. Distance-based RDA can be used with any metric or semi-metric distance measure, and it allows for partitioning components of variation in complex MANOVA models. Like RDA, it uses randomization tests to determine significance of the results. Distance-based RDA is implemented in the vegan library in R.

Summary Multivariate data contain multiple non-independent response variables measured for each replicate. Multivariate analysis of variance (MANOVA) and redundancy analysis (RDA) are the multivariate analogues of ANOVA and regression methods used in univariate analyses. Statistical tests based on MANOVA and RDA assume random, independent sampling, just as in univariate analyses, but only MANOVA requires the data conform to a multivariate normal distribution. Ordination is a set of multivariate methods designed to order samples or observations of multivariate data, and to reduce multivariate data to a smaller number of variables for additional analysis. Euclidean distance is the most natural measure of the distance between samples in multivariate space, but this distance measure can yield counterintuitive results with datasets that have many zeros, such as species abundance or presence–absence data. Other distance meas-

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Summary

ures, including Manhattan and Jaccard’s distances, have better algebraic properties, although indices of biogeographic similarity based on these measures are very sensitive to sample size effects. Principal components analysis (PCA) is one of the simplest methods for ordinating multivariate data, but it is not useful unless there are underlying correlations in the original data. PCA extracts new orthogonal variables that capture the important variation in the data and preserves the distances of the original samples in multivariate space. PCA is a special case of principal coordinates analysis (PCoA). Whereas PCA is based on Euclidean distances, PCoA can be used with any distance measure. Factor analysis is a type of reverse PCA in which measured variables are decomposed into linear combinations of underlying factors. Because the results of factor analysis are sensitive to rotation methods and because the same dataset can generate more than one set of factors, factor analysis is less useful than PCA. Correspondence analysis (CA) is an ordination tool that reveals associations between species assemblages and site characteristics. It simultaneously ordinates species and sites and may reveal groupings along both axes. However, many problems have been identified in the related detrended correspondence analysis (DCA), which was developed to deal with the horsehoe effect common in CA (and other ordination techniques). An alternative to DCA is to conduct a CA using other distance measures. Non-metric multidimensional scaling (NMDS) preserves the rank ordering of the distances, but not the distances themselves, among variables in multivariate space. Ordination methods are useful for data reduction and for revealing patterns in data. Ordination scores can be used with standard methods for hypothesis testing, as long as the assumptions of the tests are met. Classification methods are used to group objects into classes that can be identified and interpreted. Cluster analysis uses multivariate data to generate clusters of samples. Hierarchical clustering algorithms are either agglomerative (separate observations are sequentially grouped into clusters) or divisive (the entire dataset is sequentially divided into clusters). Non-hierarchical methods, such as K-means clustering, require the user to specify the number of clusters that will be created, although statistical stopping rules can be used to decide on the correct number of clusters. Discriminant analysis is a type of cluster analysis in reverse: the groups are defined a priori, and the analysis produces sample scores that provide the best classification. Jackknifing and testing of independent datasets can be used to evaluate the reliability of the discriminant function. Classification methods assume a priori that the groups or clusters are biologically relevant, whereas classic hypothesis testing would begin with a null hypothesis of random variation and no underlying groups or clusters.

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Estimation

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CHAPTER 13

The Measurement of Biodiversity

In both applied and basic ecology, much effort has gone into quantifying patterns of biodiversity and understanding the mechanisms that maintain it. Ecologists and biogeographers have been interested in species diversity at least since the writings of Humboldt (1815) and Darwin (1859), and there is a new interest from the emerging “omics” disciplines in the biodiversity of genes and proteins (Gotelli et al. 2012). Biodiversity itself encompasses a diversity of meanings (Magurran and McGill 2011), but in this chapter we focus on alpha diversity: the number of species (i.e., species richness) within a local assemblage and their relative abundances (species evenness).1 As a definition, alpha diversity may seem restrictive, but it can be applied to any collection of “objects” (e.g., individuals, DNA sequences, amino acid sequences, etc.) that can be uniquely classified into a set of exclusive “categories” (e.g., species, genes, proteins, etc.).2 By taking a small, random sample of such objects, we can apply methods from the probability calculus outlined in Chapters 1 and 2 to make inferences about the diversity of the entire assemblage.

Total diversity at the regional scale is γ diversity (gamma diversity), and the turnover or change in diversity among sites within a region is β diversity (beta diversity). The quantification of β diversity, and the contribution of α and β diversity to γ diversity have been contentious issues in ecology for over a decade. See Jost (2007), Tuomisto (2010), Anderson et al. (2011), and Chao et al. (2012) for recent reviews. 1

This simple definition can be modified to accommodate other traits of organisms. For example, all other things being equal, a community comprised of ten distantly related species, each in a different genus, would be more diverse than a community comprised of ten closely related species, all in a single genus. See Weiher (2011) and Velland et al. (2011) for recent reviews of methods of analysis for phylogenetic, functional, and trait diversity.

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Although counting the number of species in a sample might seem a simple task,3 estimating the number of species in an entire assemblage is tricky, and there are some common statistical pitfalls that should be avoided. In spite of decades of interest in the quantification of biodiversity, this is still a very active area of research, and many of the important methods described in this chapter have been developed only in the last 10 years. We first describe techniques for estimating and comparing species richness, and then consider measures of biodiversity that also incorporate species evenness. To illustrate the challenges associated with estimating biodiversity and the statistical solutions, let’s begin with an empirical example.

Estimating Species Richness A growing ecological problem is that more and more species are being transplanted (both deliberately and accidentally) beyond the limits of their historical geographic ranges and into novel habitats and assemblages. In eastern North America, the hem3

Actually, even counting (or sampling) species is not a simple task at all. Conducting a quantitative biodiversity survey is very labor-intensive and time-consuming (Lawton et al. 1998). For any particular group of organisms (e.g., ground-foraging ants, desert rodents, marine diatoms) there are specific trapping and collecting techniques—each of which has its own particular biases and quirks (e.g., bait stations, Sherman live traps, plankton tows). And once a sample has been collected, geo-referenced, preserved, cleaned up, labeled, and catalogued, each specimen has to be identified to species based on its particular morphological or genetic characters. For many tropical taxa, the majority of species encountered in a biodiversity survey may not have even been described before. Even for “well-studied” taxa in the temperate latitudes, many of the important taxonomic keys for making species identifications are buried in obscure out-of-print literature (birds are a conspicuous exception). And such keys can be difficult or impossible to use if you haven’t already been taught the basics of species identification by someone who knows how. It can take many years of practice and study to become competent and confident in the identification of even one taxonomic group. You might think that anyone calling himself or herself an ecologist would have developed this skill, but many ecologists are painfully illiterate when it comes to “reading” the biodiversity of nature and being able to identify even common species in a garden or on a campus walk. Taxonomists and museum specialists are a disappearing breed, but the great museums and natural history collections of the world still contain a treasure-trove of data and information on biodiversity—if you know how to access it. See Gotelli (2004) for an ecologist’s entrée into the world of taxonomy, and Ellison et al. (2012) for an example of how museum specimens can be compiled and analyzed to answer basic questions in ecology and biogeography.

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lock woolly adelgid (Adelges tsugae)4 is an introduced pest that is selectively killing eastern hemlock (Tsuga canadensis), a foundation species in eastern U.S. forests (Ellison et al. 2005). To study the long-term consequences of the adelgid invasion, researchers at the Harvard Forest have created large experimental plots to mimic the effects of adelgid invasion and the subsequent replacement of hemlock forests by hardwood forests (Ellison et al. 2010). In 2003, four experimental treatments were established in large (0.81 hectare) forest plots: (1) Hemlock Control (sites that had not yet been invaded by the hemlock); (2) Logged (hemlock removal); (3) Girdled (the bark and cambium of each hemlock tree in the plot were cut, which slowly kills the standing hemlock tree, just as the adelgid does); and (4) Hardwood Control (the forest type expected to replace hemlock stands after the adelgid invasion). In 2008, invertebrates were censused in the four plot types by using an equal number of pitfall traps in each treatment (Sackett et al. 2011). Table 13.1 summarizes the data for spiders collected in the four treatments. Although 58 species were collected in total, the number of species in each treatment ranged from 23 (Hemlock Control) to 37 (Logged). Does the number of spider species differ significantly among the four treatments? Although it would seem that more spider species were collected in the Logged treatment, we note that the Logged treatment also had the greatest number of individuals (252). The Hemlock Control had only 23 species, but perhaps that is not surprising, considering that only 106 individual spiders were collected. In fact, the rank order of species richness among the four treatments follows exactly the rank order of spider abundance (compare the last two rows of Table 13.1)! If you are surprised by the fact that more individuals and species of spiders were collected in the Logged versus the Hemlock Control plots, think again. Logging generates a lot of coarse and fine woody debris, which is ideal microhabitat for many spiders. Moreover, the elimination of the cool shade provided by the hemlock trees boosts the air and soil temperatures in the plot (Lustenhouwer et al. 2012). This 4

The hemlock woolly adelgid is native to East Asia. This small phloem-feeding insect was introduced to the United States in 1951 on nursery stock—Asian hemlock saplings destined for landscaping and garden stores—imported into Richmond, Virginia. It was next collected in the mid-1960s in the Philadelphia area, but did not attract any attention until the early 1980s, when its populations irrupted in Pennsylvania and Connecticut. The adelgid subsequently spread both north and south, leaving vast stands of dead and dying hemlocks in its wake (Fitzpatrick et al. 2012). In Northeast North America, the spread of the adelgid is limited by cold temperatures, but as regional climates warm, the adelgid continues to move northward. Although individual trees can be treated with insecticide to kill the adelgid, chemical control is impractical on a large scale. Biological control is being explored (Onken and Reardon 2011), but with limited success so far.

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TABLE 13.1 Spider diversity data Hemlock Control

Species

Girdled

Hardwood Control

Logged

Agelenopsis utahana

Agroeca ornata

Callobius bennetti

Castianeira longipalpa

Centromerus cornupalpis

Centromerus persolutus

Ceraticelus minutus

Ceratinella brunnea

Cicurina arcuata

Amaurobius borealis

Pardosa moesta

Zelotes fratris

Total number of individuals

106

168

250

252

Total number of species

Counts of the number of individuals of 58 spider species collected from four experimental treatment groups in the Harvard Forest Hemlock Removal Experiment. Data from Sackett et al. (2011); see the online resources for the complete data table.

matters because spiders (and all invertebrates) are ectotherms—that is, their body temperature is not internally regulated, as in so-called “warm-blooded” birds and mammals. Thus, the population sizes of ectotherms often will increase after logging or other disturbances because the extra warmth afforded by the elimination of shade gives individuals more hours in the day and more days in the year during which they can actively forage. Finally, the increase in temperature may increase the number of spiders collected even if their population sizes have not increased: spiders are more active in warmer temperatures, and their extra movement increases the chances they will be sampled by a pitfall trap. For all these reasons, we must make some kind of adjustment to our estimate of spider species richness to account for differences in the number of individuals collected.

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You also should recognize that the sample plots are large, the number of invertebrates in them is vast, and the counts in this biodiversity sample represent only a tiny fraction of what is out there. From the perspective of a 5-mm long spider, a 0.81-ha plot is substantially larger than all five boroughs of New York City are to an individual human! In each of the plots, if we had collected more individual spiders, we surely would have accumulated more species, including species that have so far not been detected in any of the plots. This sampling effect is very strong, and it is pervasive in virtually all biodiversity surveys (whether it is recognized or not). Even when standard sampling procedures are used, as in this case, the number of individuals in different collections almost never will be the same, for both biological and statistical reasons (Gotelli and Colwell 2001). One tempting solution to adjust for the sampling effect would be to simply divide the number of species observed by the number of individuals sampled. For the Logged treatment, this diversity estimator would be 37/252 = 0.15 species/individual. For the Hemlock Control treatment, this diversity estimator would be 23/106 = 0.22 species/individual. Although this procedure might seem reasonable, the sampling curve of species richness (and other diversity measures) is distinctly non-linear in shape. As a consequence, a simple algebraic rescaling (such as dividing species richness by sampling effort or area) will substantially over-estimate the actual diversity in large plots and should always be avoided. Standardizing Diversity Comparisons through Random Subsampling

How can we validly compare species richness in the Logged plots versus the Hemlock Control plots, when there are 252 individual spiders sampled from the Logged plots, but only 106 sampled from the Hemlock Control plots? An intuitively appealing approach is to use a random subsample of the individuals from the larger sample, drawing out the same number of individuals as were found in the smaller sample. As an analogy (following Longino et al. 2002), imagine that each individual spider in the Logged plot is a single jelly bean. The jelly beans are of 37 colors, each one corresponding to a particular species of spider in the sample from the Logged treatment. Put all 252 jelly beans in a jar, mix them thoroughly, and then draw out exactly 106 jelly beans. Count the number of colors (i.e., species) in this random subsample, and then compare it to the observed number of species in the sample from the Hemlock Control treatment. Such “candy-jar sampling”—known as rarefaction—allows for a direct comparison of the number of species in the Hemlock Control and Logged plots based on an equivalent number of individuals in both samples (106). This method of drawing random subsamples can be applied to any number of individuals less than or equal to the original sample size. To rarefy a sample in this manner is

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to make it less dense, and a rarefaction curve gives the expected number of species and its variance for random samples of differing sizes.5 Table 13.2 (see next page) shows the results of a computer simulation of this exercise. This particular subsample of 106 individuals from the Logged sample contains 24 species, which is pretty close to the 23 species observed in the Hemlock Control sample. Of course, if we were to replace all the jelly beans and repeat this exercise, we would get a slightly different answer, depending on the mix of individuals that are randomly drawn and the species they represent.6 Figure 13.1 shows a histogram of the counts of species richness from 1000 such random draws. For this particular set of random draws, the simulated species richness ranges from 19 to 32, with a mean of 26.3. The observed richness of 5

Rarefaction has an interesting history in ecology. In the 1920s and then again in the 1940s, several European biogeographers independently developed the method to compare taxonomic diversity ratios, such as the number of species per genus (S/G; Järvinen 1982). In his influential 1964 book, Patterns in the balance of nature, C. B. Williams argued that most of the observed variation in S/G ratios reflected sampling variation (Williams 1964). The early controversy over the analysis of S/G ratios foreshadowed a much larger conflict in ecology over the use of null models in the 1970s and 1980s (Gotelli and Graves 1996). In 1968, the marine ecologist Howard Sanders proposed rarefaction as a method for standardizing samples and comparing species diversity among habitats (Sanders 1968). Sanders’ mathematical formula was incorrect, but his notion that sample size adjustments must be made to compare diversity was sound. Since then, the original individual-based rarefaction equation has been derived many times (Hurlbert 1971), as has the equation for sample-based rarefaction (Chiarucchi et al. 2008), and there continue to be important statistical developments in this literature (Colwell et al. 2012).

Note that for a single simulation, the 106 individual jelly beans are randomly sampled without replacement. Drawing cards from a deck is an example of sampling without replacement. For the first card drawn, the probability of obtaining a particular suit (clubs, diamonds, hearts, or spades), is 13/52 = 1/4. However, these probabilities change once the first card is drawn, and they are conditional on the result of the first draw. For example, if the first card drawn is a spade, the probability that the second card drawn will also be a spade is 12/51. If the first card drawn is not a spade, the probability that the second card drawn is a spade is now 13/51. However, in the rarefaction example, all of the 106 jelly beans would be replaced before the start of the next simulation, sampling with replacement. A familiar example of sampling with replacement is the repeated tossing of a single coin. Sampling with or without replacement correspond to different statistical distributions and different formulae for the expectation and the variance. Most probability models (as in Chapters 1 and 2) assume sampling with replacement, which simplifies the calculations a bit. In practice, as the size of the sampling universe increases relative to the size of the sample, the expected values converge for the two models: if the jelly bean jar is large enough, the chances of getting a particular color are nearly unchanged whether or not you replace each jelly bean before you draw another one.

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Figure 13.1 Histogram of species richness counts for

455

200

1000 random subsamples of 106 individuals from the Logged plots of the Harvard Forest Hemlock Removal Experiment (see data in Table 13.1). The blue line is the observed species richness (23) from the 106 individuals in the Hemlock Control plots, and the dashed lines bracket the 95% confidence interval for the simulated distribution.

Frequency

150

100

0 15

20 25 30 35 Simulated species richness

23 species for the Hemlock Control treatment is on the low side (P = 0.092 for the lower tail of the distribution), but still falls within the 95% confidence interval of the simulated distribution (22 to 31 species). We would conclude from this analysis that the Hemlock Control and Logged treatments do not differ significantly in the number of species they support. Rarefaction Curves: Interpolating Species Richness

The full rarefaction curve for the spiders of the Logged treatment, as shown in Figure 13.2, illustrates the general features characteristic of any individual-based rarefaction curve. We call this kind of rarefaction curve “individual-based” because the individual organism is the unit of sampling. The x-axis for the graph of an individual-based rarefaction curve is the number of individuals, and the

Species richness

Figure 13.2 Individual-based rarefaction curve for the 10

100 150 200 Number of individuals

250

spider data from the Logged plots in the Harvard Forest Hemlock Removal Experiment (see data in Table 13.1). The simulation is based on 1000 random draws at every sampling level from 1 to 252. The black line is the average of the simulated values, and the blue envelope is the parametric 95% confidence interval. The point represents the original reference sample (252 individuals, 37 species).

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TABLE 13.2 Observed and randomly subsampled counts of spiders

Logged

Logged (random subsample)

Hemlock Control

Agelenopsis utahana Agroeca ornata Amaurobius borealis Callobius bennetti Castianeira longipalpa Centromerus cornupalpis Centromerus persolutus Ceraticelus minutus

1 10 22 3 3 0 0 1

0 3 10 0 2 0 0 0

1 2 27 4 0 0 1 1

Ceratinella brunnea Cicurina arcuata Cicurina brevis Cicurina pallida Cicurina robusta Collinsia oxypaederotipus Coras juvenilis Cryphoeca montana Dictyna minuta

4 1 0 0 1 2 0 2 0

2 0 0 0 0 1 0 1 0

3 0 0 0 0 0 1 2 0

0 0 0 4 0 2 0 0 0 8 1 1 15 1 0 1

0 0 0 3 0 1 0 0 0 3 0 0 7 1 0 0

0 1 1 0 0 0 0 1 1 2 0 0 11 0 0 0

Species

Emblyna sublata Eperigone brevidentata Eperigone maculata Habronattus viridipes Helophora insignis Hogna frondicola Linyphiid sp. 1 Linyphiid sp. 2 Linyphiid sp. 5 Meioneta simplex Microneta viaria Naphrys pulex Neoantistea magna Neon nelli Ozyptila distans Pardosa distincta

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TABLE 13.2 Continued

Species

Logged

Pardosa moesta Pardosa xerampelina

Logged (random subsample)

Hemlock Control

Pelegrina proterva

Phidippus whitmani

Phrurotimpus alarius

Phrurotimpus borealis

Pocadicnemis americana

Robertus riparius

Scylaceus pallidus

Tapinocyba minuta

Tapinocyba simplex

Tenuiphantes sabulosus

Tenuiphantes zebra

Trochosa terricola

Unknown morphospecies 1

Wadotes calcaratus

Wadotes hybridus

Walckenaeria digitata

Walckenaeria directa

Walckenaeria pallida

Xysticus elegans

Xysticus fraternus

Zelotes duplex

Zelotes fratris

252

106

Pirata montanus

Total number of individuals Total number of species

The first and third columns give the counts of spiders from the Logged and Hemlock Control treatments. The second column gives the counts of spiders from a computer-generated random subset of 106 individuals from the Logged sample. By rarefying the data from the Logged treatment, direct comparisons can be made to observed species richness in the Hemlock Control treatment based on a standardized number of individuals (106).

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y-axis is the number of species. At the low end, the individual-based rarefaction begins at the point (1,1) because a random draw of exactly 1 individual always yields exactly 1 species. With two individuals drawn, the observed species richness will be either 1 or 2, and the expected value for two individuals is thus an average that is between 1.0 and 2.0. The smoothed curve of expected values continues to rise as more individuals are sampled, and it does so with a characteristic shape: it is always steepest at the low end, with the slope becoming progressively shallower as sampling increases. Such a shape results because the most common species in the assemblage are usually picked up in the first few draws. With additional sampling, the curve continues to rise, but more slowly, because the remaining unsampled species are, on average, progressively less common. In other words, most of the new individuals sampled at that stage represent species that have already been added. Because most of the individuals in the collection are included in larger random draws, the expected species number eventually rises to its highest point, indicated by the dot in Figure 13.2. The high point represents a sample of the same number of individuals as the original collection. Note that this end-point of species richness does not represent the true asymptote of richness for the entire assemblage—it represents only the number of species that were present in the original sample. If the sample had been larger to begin with, it probably would have contained more species. Of course, if empirical sampling is intensive enough, all of the species in an assemblage will have been encountered, and the sampling curve will reach a flat asymptote. In practice, the asymptote is almost never reached, because a huge amount of additional sampling is usually needed to find all of the rare species. Rarefaction is thus a form of interpolation (see Chapter 9): we begin with the observed data and rarefy down to progressively smaller sample sizes. The variance in Figure 13.2 also has a characteristic shape: it is zero at the low end because all random samples of 1 individual will always contain only 1 species. It is also zero at the high end because all random samples of all individuals will always yield exactly the observed species richness of the original sample. Between these extremes, the variance reflects the uncertainty in species number associated with a particular number of individuals (see Figure 13.1). When estimated by random subsampling, the variance is conditional on the empirical sample and will always equal zero when the size of the subsample equals that of the original data. We will return to this variance calculation later in the next section. Figure 13.3 again illustrates the rarefaction curve for the Logged treatment (and its 95% confidence interval), but with the added rarefaction curves for the other three treatments. It is easy to see that the Hemlock Control rarefaction

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Figure 13.3 Individual-based rarefaction curves for the four experimental treatments in the Harvard Forest Hemlock Removal Experiment. The lines are the expected species richness based on 1000 random draws from the data for each plot (see Table 13.1). Black: Logged plots; light gray: Hardwood Control plots; dark gray: Girdled plots; blue: Hemlock Control plots. The blue area is the parametric 95% confidence interval for the Logged plots.

30 Species richness

459

100 150 200 Number of individuals

250

curve lies within the 95% confidence interval of the Logged rarefaction curve. The Hardwood Control rarefaction curve has the lowest species richness for a given sampling level, whereas the Girdled rarefaction curve lies very close to the Hemlock Control curve. Notice how all the curves converge and even cross at the lower end. When rarefaction curves cross, the ranking of species richness will depend on what sampling level is chosen. The Expectation of the Individual-Based Rarefaction Curve

Formally, suppose an assemblage consists of N total individuals, each belonging to one of S different species. Species i has Ni individuals, which represents a proportion of the total pi = Ni /N. By definition, these pi values add up to 1.0 when summed across all species: ∑ Si =1 pi = 1.0 . We stress that N, S, and pi represent the complete or “true” values for an assemblage: if we only knew what these numbers were, we would not have to worry about sampling effects, or even about sampling! We could just compare these numbers directly. Instead, we have only a small representative sample of data from this assemblage. Specifically, we have a reference sample of only n individuals (where n is a number usually much smaller than N), with Sobs species counted, and Xi individuals of each species i. Because the sampling is incomplete, some species in the assemblage will be missing from the reference collection. Thus, Xi = 0 for some values of i. The estimated proportion of abundance represented by each species i is pˆ i =Xi /n. With individual-based rarefaction, we would like to estimate the expected number of species in a small subset of m individuals drawn from our reference sample of n. If we knew the underlying pi values, we could derive the expected

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number of species using some simple laws of probability (see Chapter 2). First, if we randomly draw out m individuals, with a multinomial sampling model7 the probability of not encountering species i in any of the m trials is: p(not sampling species i) = (1 – pi)m

(13.1)

Therefore, the probability of capturing species i at least once with m trials is: p(sampling species i) = 1 – (1 – pi)m

(13.2)

Summing these probabilities across all i species gives the expected number of species in a sample of m individuals: s

i =1

m m Sind (m ) = ∑ ⎡1 − (1 − pi ) ⎤ = S − ∑ (1 − pi ) ⎣ ⎦

(13.3)

Note that the larger m is, the smaller the term within the summation sign, and therefore the closer Sind(m) is to the complete S. For the empirical reference sample of n individuals, an unbiased estimator of the expected number of species from a small subsample of m individuals comes from the hypergeometric distribution:8 7

The multinomial probability distribution generalizes the binomial distribution that was explained in detail in Chapter 2. Recall that a binomial distribution describes the numbers of successes of a set of n independent Bernoulli trials: experiments that have only two possible outcomes (present or absent, reproduce or not, etc.). If there are many possible (k) discrete outcomes (red, green, blue; male, female, hermaphrodite; 1, 2, 3, …, many species) and each trial results in only one of these possible outcomes, each with probability pi (i = 1, 2, …, k), and there are n trials, then the random variable Xi has a multinomial distribution with parameters n and p. The probability distribution function for a multinomial random variable is P(X ) =

n! p1x1 ⋅… ⋅ pkxk x1 !⋅… ⋅ xk !

the expected value of any outcome E{Xi } = npi , the variance of an outcome V(Xi ) = npi (1 – pi ), and the covariance between two distinct outcomes (i ≠ j) is Cov(Xi , Xj ) = –npi p j . 8

Like the binomial distribution, the hypergeometric distribution is a discrete probability distribution that models the probability of k successes in n draws from a population of size N with m successes. The difference between the two is that draws or trials in a

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Sind (m ) = Sobs − ∑ ⎡ ⎣ x i >0

( ) / ( )⎤⎦ n− X i m

n m

(13.4)

Equation 13.4 gives the exact solution for the expected number of species, sampling m individuals without replacement from the original collection of n individuals. In practice, a very similar answer usually results from Equation 13.3, sampling with replacement and using Xi/n as an estimator for pi. Both equations also can be estimated by the simple random sampling protocol that was used to generate Table 13.2 and Figures 13.1 and 13.2. Heck et al. (1975) give an equation for the variance of Equation 13.4, but we don’t need to use it here. However, we should note that the variance for standard rarefaction is conditional on the collection of n individuals: as the subsample m approaches the reference sample n in size, this variance and the width of the associated 95% confidence interval approaches zero (see Figure 13.2). Sample-Based Rarefaction Curves: Massachusetts Ants

One difficulty with the “jelly bean” sampling model is that it assumes that the individual organism is the unit that is randomly sampled. However, most ecological sampling schemes use some larger collecting unit, such as a quadrat, trap, plot, transect, bait, or other device that samples multiple individuals. For example, the spider data compiled in Table 13.1 actually were pooled from two plots per treatment, with four pitfall traps per plot. It is these sampling units that represent statistically independent replicates, not the individuals themselves (see Chapters 6 and 7).

hypergeometric distribution are sampled without replacement, whereas draws in a binomial distribution are sampled with replacement. The probability distribution function of a hypergeometric random variable is P(X ) =

( )( ) ( ) m k

N −m n− k

N n

()

where ba is the binomial coefficient (see Chapter 2). The expected value of a hypergeometric random variable is m E(X ) = n N and its variance is ⎛ m ⎞ ⎛ N − m⎞ ⎛ N − n⎞ Var ( X ) = ⎜ n ⎟ ⎜ ⎝ N ⎠ ⎝ N ⎟⎠ ⎜⎝ N − 1 ⎟⎠

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In some kinds of studies, it may not even be possible to count individual organisms within a sample. For example, many marine invertebrates, such as corals and sponges, as well as many perennial plants, grow as asexual clones, making it difficult to define an “individual.” In such cases, we can only score the presence or incidence of a species in a sample; we cannot count its abundance. But whether the individuals are counted or not, there are multiple sampling units for each treatment or habitat that is surveyed. We therefore apply sample-based rarefaction to such data sets. For example, Table 13.3 gives the number of ant nests of different species that were detected within 12 standardized sample plots (Allen’s Pond through Peaked Mountain) in Massachusetts cultural grasslands.9 Each row is a different ant species, and each column is a different sample plot in a grassland habitat. In this data set, the entry is the number of ant nests of a particular species detected in a particular sample plot.10 The data are organized as an incidence matrix, in which each entry is simply the presence (1) or absence (0) of a species in a sample. Incidence matrices are most often used for sample-based rarefaction. We have comparable data for 11 other plots from oak-hickory-white pine forests, and five plots from successional shrublands. In the spider example, we used individual-based rarefaction to compare species richness among four experimental treatments. In this analysis, we use 9

A few words on the origin of the phrase “cultural grasslands.” What are called “natural” grasslands in New England are fragments of a coastal habitat that was more widespread during the relatively warm Holocene climatic optimum, which occurred 9000–5000 years ago. However, most contemporary grasslands in New England are rather different: they are the result of clearing, agriculture, and fire, and have been maintained for many centuries by the activities of Native Americans and European settlers. These cultural grasslands currently support unique assemblages of plants and animals, many of which do not occur in more typical forested habitats. Motzkin and Foster (2002) discuss the unique history of cultural grasslands and their conservation value in New England. 10

The ant nests were discovered by standardized hand searching at each site for one person-hour within a plot of 5625 m2. Hand-searching for nests is a bit of an unusual survey method for ants, which are usually counted at baits or pitfall traps. Although we can count individual worker, queen, or male ants, a quirky feature of ant biology complicates the use of these simple counts for biodiversity estimation. Ant workers originate from a nest that is under the control of one or more queens. The nest is in effect a “super-organism,” and it is the nest, not the individual ant worker, that is the proper sampling unit for ants. When dozens of ant workers of the same species show up in a pitfall trap, it is often because they all came from a single nest nearby. Thus, counting individual ant workers in lieu of counting ant nests would be like counting individual leaves from the forest floor in lieu of counting individual trees. Therefore, pitfall data for ants are best analyzed with sample-based rarefaction. Gotelli et al. (2011) discuss some of the other challenges in the sampling and statistical analysis of ant diversity. The unique biological features of different plants and animals often dictate the way that we sample them, and constrain the kinds of statistical analyses we can use.

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Estimating Species Richness TABLE 13.3 Data for sample-based rarefaction

Species

Allen’s Pond

Boston Nature Center

Brooks Woodland

Daniel Webster

Doyle Center

Drumlin Farm

Elm Hill

Graves Farm

Moose Hill

Nashoba Brook

Old Town Hill

Peaked Mountain

Sites

Aphaenogaster rudis complex

Brachymyrmex depilis

Formica incerta

Formica lasiodes

Formica neogagates

Formica neorufibarbis

Formica pergandei

Formica subsericea

Lasius alienus

Lasius flavus

Lasius neoniger

0 15

1 12

3 11

Lasius umbratus

Myrmica americana

Myrmica detritinodis

0 12

Myrmica nearctica

Myrmica punctiventris

Myrmica rubra

Ponera pennsylvanica

Prenolepis imparis

Solenopsis molesta

Stenamma brevicorne

Stenamma impar

Tapinoma sessile

Temnothorax ambiguus

Temnothorax curvispinosus

Tetramorium caespitum

0 13

Each row is a species, each column is a sampled cultural grassland site, and each cell entry is the number of ant nests recorded for a particular species in a particular site. Although this data set contains information on abundance, the same analysis could be carried out with an incidence matrix, in which each cell entry is either a 0 (species is absent) or a 1 (species is present). The data for the other two habitats—the oak-hickory-white pine forest and the successional shrublands—are available in the book’s online resources.

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Figure 13.4 Sample-based rarefaction of num25 20 Species richness

ber of ant species as a function of the number of sample plots surveyed in Massachusetts. Black: oak-hickory-white pine forests; blue: cultural grasslands (see data from Table 13.3); gray: successional shrublands. The blue band is the 95% confidence interval for the oak-hickory-white pine forest.

15 10 5

6 8 10 Number of samples

sample-based rarefaction to compare ant species richness among three habitat types. The principle is exactly the same, but instead of randomly sampling individuals, we randomly sample entire plots. Figure 13.4 illustrates the samplebased rarefaction curves for the three habitat types. As with individual-based rarefaction, the sample-based rarefaction curve rises steeply at first, as common species are encountered in the first few plots, and then more slowly, as rare species are encountered as more plots are included. The endpoint represents the number of species and samples in the original reference sample. In this example, the reference samples had plot numbers of 5 (successional shrublands), 11 (oakhickory-white pine forests) and 12 (cultural grasslands). The variance (and a 95% confidence interval) also can be constructed from the random samples. As before, the variance is zero at the (maximum) reference sample size because, when all of the plots are used, the number of species is exactly the same as in the original data set. However, the variance at the low end does not look like that of the individual-based rarefaction curve. In the individual-based rarefaction curve, a subsample of one individual always yields exactly one species. But in the sample-based rarefaction curve, a subsample of one plot may yield more than one species, and there is uncertainty in this number depending on which plot is randomly selected. Therefore, the sample-based rarefaction curve usually has more than one species at its minimum of one sample, and the variance associated with that estimate is greater than zero. As in the previous discussion of individual-based rarefaction, this variance estimator for sample-based rarefaction is conditional on the particular samples in hand. Sample-based rarefaction curves effectively control for differences in the number of samples collected in the different habitats being compared. However, there may still be differences in the number of individuals per sample that can affect the species richness estimate. Because this data set has counts of individual ant nests in

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Estimating Species Richness

465

Figure 13.5 Sample-based rarefaction, rescaled to the average number of individual ant nests per sample on the x-axis. These are rarefaction curves of the same ant data used to generate Figure 13.4, but the x-axis now illustrates the average number of individuals per sample, not the number of samples. Note that the rank ordering of the rarefaction curves is different in the two graphs. Black: oak-hickory-white pine forests; blue: cultural grasslands (see data from Table 13.3); gray: successional shrublands.

Species richness

20 15 10 5

100 150 200 250 300 Number of individuals

350

each plot, we can take one further step, which is to re-plot the sample-based rarefaction curves (and their variances) against the average number of individuals (or incidences) per plot.11 This potentially shifts and stretches each rarefaction curve depending on the abundance (or incidence) per sample (Gotelli and Colwell 2001). For the Massachusetts ant data, the rank order of the sample-based rarefaction curves (see Figure 13.4) changes when they are plotted against abundance, as in Figure 13.5. Specifically, the cultural grassland curve now lies above the other two (it is barely within the confidence interval for the oak-hickory-white pine forest curve). This example illustrates that the estimate of species richness depends not only on the shape of the rarefaction curve, but also on the number of samples and the number of individuals or incidences per sample being compared. Species Richness versus Species Density

Although hundreds of ecological papers analyze species richness, it is actually better to refer to the measured number of species as species density, the number of species per sample unit (James and Wamer 1982). As illustrated by individual-based rarefaction, species density depends on two components: Species Individuals Species = × Sample Sample Individual

(13.5)

Two communities might differ in the number of species / sample because of either differences in the number of species / individual (which is quantified with 11

With an incidence matrix, expected species richness can be re-plotted against the average number of incidences (species occurrences) per sample. However, incidences sometimes can be difficult to interpret because they may not correspond to a discrete “individual” or other sampling unit that contains biodiversity information.

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the rarefaction curve) or differences in the number of individuals / sample. Variation in the number of individuals / sample may reflect differences in sampling effort (how many individuals were collected) or detection probability (e.g., pitfall traps catch more samples on warm days than on cold days; see Chapter 14), or differences in biological factors (e.g., gradients in productivity or available energy). Rarefaction is a straightforward way to control for differences in the number of individuals per sample and their effect on species richness. For a sample-based data set, in which there are several plots per sample, and abundance is measured within each plot, the relationship is: Species Plots Individuals Species = × × Sample Sample Plot Individual

(13.6)

As we noted earlier, it is never appropriate to estimate diversity by computing these values as simple ratios, but Equations 13.5 and 13.6 do illustrate how the sampling properties of the data contribute to the observed species density.

The Statistical Comparison of Rarefaction Curves This chapter (along with Chapter 14) emphasizes the methods for the proper estimation of diversity. Once rarefaction curves have been calculated, species richness estimated for any level of sampling can be compared using conventional statistical methods, including regression (see Chapter 9) and analysis of variance (see Chapter 10). Hypothesis testing with rarefaction curves is possible, although there has not been much published work in this area. To test whether species richness differs among rarefaction curves for a particular level of sampling, a simple, conservative test is to examine whether there is any overlap in the 95% confidence intervals calculated for each curve. Payton et al. (2003) recommend a less conservative approach with approximate 84% confidence intervals to control for Type I error. However, both methods assume equal variances in the two groups. The results will depend on the sampling level that is chosen for comparison (particularly if the rarefaction curves cross), and on the way in which the variance of the rarefaction curve is calculated (see next section).12 12 What is needed is a general test for differences among rarefaction curves. In many ecological studies, the implicit null hypothesis is that the two samples differ no more in species richness than would be expected if they were both drawn randomly from the same assemblage. However, in many biogeographic studies, the null hypothesis is that the samples differ no more in species richness than would be expected if the underlying shape of the species abundance distribution were the same in both assemblages. For many biogeographic comparisons, there may be no shared species among the regions, but it is still of

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The Statistical Comparison of Rarefaction Curves

Assumptions of Rarefaction

Whether using sample-based or individual-based rarefaction, the following assumptions should be considered: As illustrated in Figures 13.3 and 13.4, rarefaction curves converge at small sample sizes, and the differences among curves can become highly compressed. Because all rarefaction curves have to be compared at the sampling intensity of the smallest reference sample, it is important that this sample be of sufficient size for comparison to the others.13 There is no set amount, but in practice, rarefaction curves based on reference samples of fewer than 20 individuals or five samples usually are inadequate. SUFFICIENT SAMPLE SIZE

As discussed earlier in this chapter (see Footnote 3), all sampling methods have their biases, such that some species will be overrepresented and others will be underrepresented or even missing. There is no such thing as a truly “random” sampling method that is completely unbiased. For this reason, it is important that identical sampling methods are used for all comparisons. If quadrat surveys are used in grasslands, but point counts are used

COMPARABLE SAMPLING METHODS

interest to ask whether they differ in species richness. We are starting to explore some bootstrapping analyses in which we measure the expected overlap among rarefaction confidence intervals calculated for different curves. Stay tuned! 13 Alroy (2010), Jost (2010), and Chao and Jost (in press) recently have suggested a new approach to the standardization of diversity curves. Rather than comparing them at a constant number of samples or individuals, rarefaction curves can be compared at a common level of coverage or completeness. Coverage is the percentage of the total abundance of the assemblage that is represented by the species in the reference sample. Thus, a coverage of 80% means that the species represented in the reference sample constitute 80% of the abundance in the assemblage. The abundance of the undetected species in the reference sample constitutes the remaining 20%. A good estimator of coverage comes from the work of Alan Turing, a British computer scientist who made fundamental contributions to statistics (see Footnote 21 in this chapter for more details on Turing). Turing’s simple formula is Coverage ≈ 1.0 – f1/n, where f1 is the number of species represented by exactly 1 individual (i.e., the number of singletons), and n is the number of individuals in the reference sample (see the section Asymptotic Estimators: Extrapolating Species Richness for more on singletons). Coverage-based rarefaction standardizes the data to a constant level of completeness, which is not exactly the same as standardizing to a common number of individuals or samples. However, rarefaction curves based on coverage have the same rank order as rarefaction curves based on the number of individuals or the number of samples, so we still advocate the use of traditional rarefaction for ease of interpretation. See Chao and Jost (in press) for more details on coverage-based rarefaction.

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in wetlands, the comparison of diversity in the two habitats will be confounded by differences in the sampling methods.14 The implicit null hypothesis in many ecological applications of rarefaction is that the samples for different habitats or treatments have been drawn from a single underlying assemblage. In such a case, the samples will often have many shared species, as in the spider data of Table 13.1. Thus, some degree of taxonomic similarity often is listed as an assumption of rarefaction (Tipper 1979). However, in biogeographic analyses (such as comparisons of Asian versus North American tree species richness), there may be few or no species shared among the comparison groups. For either biogeographic or ecological comparisons, rarefaction curves reflect only the level of sampling and the underlying relative abundance distribution of the component species. Rarefaction curves often do not reflect differences among assemblages in species composition,15 and there are more powerful, explicit tests for differences in species composition among samples (Chao et al. 2005). TAXONOMIC SIMILARITIES OF SAMPLES

The jelly bean analogy for sampling assumes that the assemblage meets the closure assumption, or that the assemblage at least is circ*mscribed enough so that individuals sampled from

CLOSED COMMUNITIES OF DISCRETE INDIVIDUALS

Researchers sometimes combine the data from different survey methods in order to maximize the number of species found. Such a “structured inventory” is valid as long as the same suite of methods is used in all the habitats that are being compared (Longino and Colwell 1997). It is true that some species may show up only in one particular kind of trap. However, such species are often rare, and it is hard to tell whether they are only detectable with one kind of trapping, or whether they might show up in other kinds of traps if the sampling intensity were increased. 15

The spider data in Table 13.2 illustrate this principle nicely. Although the rarefaction curves for the Logged and Hemlock Control treatments are very similar (see Figure 13.3), there are nevertheless differences in species composition that cannot be explained by random sampling. These differences in species composition are best illustrated with columns 2 and 3 of Table 13.2, which show the complete data for the Hemlock Control treatment (column 3, n = 106 individuals, 23 species) and a single rarefied sample of the data for the Logged treatment (column 2, n = 106 individuals, 24 species). The rarefied sample for the Logged treatment contained 40 individuals of Pardosa xerampelina, whereas zero individuals of this species were observed in the Hemlock Control treatment. Conversely, the Hemlock Control treatment contained 20 individuals of Pirata montanus whereas the rarefied sample from the Logged treatment contained only five individuals of this species. Even though total species richness was similar in the two habitats, the composition and species identity clearly were different.

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The Statistical Comparison of Rarefaction Curves

this assemblage can be counted and identified. If individuals cannot be readily counted, then a sample-based rarefaction design is needed.16 Rarefaction directly addresses the issue of undersampling, and the fact that there may be many undetected species in an assemblage. However, the model assumes that the probability of detection or capture per individual is the same among species and among comparison groups (habitats or treatments; see Chapter 14 for formal statistical models of detection probability). Consequently, any differences in the detection or capture probability per species reflect underlying differences in commonness and rarity of different species.17

CONSTANT DETECTION PROBABILITY PER INDIVIDUAL

Individual-based rarefaction assumes that individuals of the different species are well mixed and occur randomly in space. However, this is often not true, and the more typical pattern is that individuals of a single species are clumped or spatially aggregated (see the discussion of the coefficient of dispersion in Chapter 3). In such cases, the spatial scale of sampling needs to be increased to avoid small-scale patchiness (see Chapter 6). Alternatively, use sample-based rarefaction.18 SPATIAL RANDOMNESS OF INDIVIDUALS

INDEPENDENT, RANDOM SAMPLING

As with all of the statistical methods described in this book, the individuals or samples should be collected randomly and independently. Both rarefaction methods assume that the sampling itself does not

The assumption of a closed community can be difficult to justify if the assemblage is heavily affected by migration from adjacent habitats (Coddington et al. 2009), or if there is temporal variation in community structure from ongoing habitat change (Magurran 2011). In many real communities, the jelly bean jar probably “leaks” in space and time. 17

Hierarchical sampling models (Royle and Dorazio 2008) can be used to explicitly model the distinct components of the probability of occupancy and the probability of detection, given that a site is occupied. These models use either maximum likelihood or Bayesian approaches to estimate both probabilities and the effects of measured covariates. Chapter 14 introduces these models, which can be applied to many problems, including the estimation of species richness (Kéry and Royle 2008). 18

You may be a tempted to pool the individuals from multiple samples and use individual-based rarefaction. However, if there is spatial clumping in species occurrences, the individual-based rarefaction curve derived from the pooled sample will consistently over-estimate species richness compared to the sample-based rarefaction curve. This is yet another reason to use sample-based rarefaction: it preserves the small-scale heterogeneity and spatial clumping that is inherent in biodiversity samples (Colwell et al. 2004).

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affect the relative abundance of species, which is statistically equivalent to sampling with replacement (see Footnote 6 in this chapter). For most assemblages, the size of an ecological reference sample is very small compared to the size of the entire assemblage, so this assumption is easy to meet even when the individuals are not replaced during sampling.

Asymptotic Estimators: Extrapolating Species Richness Rarefaction is an effective method of interpolating and estimating species richness within the range of observed sampling effort. However, with continued sampling of a closed assemblage, the species accumulation curve would eventually level out at an asymptote.19 Once the asymptote is reached, no new species would be added with additional sampling. The asymptote of S species with N individuals thus represents the total diversity of the assemblage. In contrast, the reference sample consists of only S observed species (Sobs) with n individuals. Unless the reference sample is very large or there are very few species in the assemblage, Sobs 0 and f2 >0):23 s 2Chao1

⎡1 ⎛ f ⎞2 ⎛ f ⎞3 1 ⎛ f ⎞4⎤ = f2 ⎢ ⎜ 1 ⎟ + ⎜ 1 ⎟ + ⎜ 1 ⎟ ⎥ 4 ⎝ f2 ⎠ ⎥ ⎝ f2 ⎠ ⎢⎣ 2 ⎝ f 2 ⎠ ⎦

(13.9)

For the Hemlock Control treatment, the estimated variance is 34.6, with a corresponding 95% confidence interval of 18.2 to 41.3 species. For sample-based rarefaction, the formulas are similar, but instead of using f1 and f2 for the number of singletons and doubletons, we use q1, the number of uniques (= species occurring in exactly 1 sample) and q2, the number of duplicates (species occurring in exactly 2 samples). The formula for Chao2, the expected number of species with sample-based data, also includes a small bias correction for R, the number of samples in the data set: 2 ⎛ R − 1 ⎞ q1 Chao2 = Sobs + ⎜ if q2 > 0 ⎝ R ⎟⎠ 2q2

(13.10)

⎛ R − 1 ⎞ q1 ( q1 − 1) Chao2 = Sobs + ⎜ if q2 = 0 ⎝ R ⎟⎠ 2 ( q2 + 1)

(13.11)

The corresponding variance estimate for sample-based incidence data (with q1 > 0 and q2 > 0) is: s 2Chao2

3 4 ⎡ A⎛ q ⎞2 A 2 ⎛ q1 ⎞ ⎤ 2 ⎛ q1 ⎞ 1 ⎥ = q2 ⎢ ⎜ ⎟ + A ⎜ ⎟ + 4 ⎜⎝ q2 ⎟⎠ ⎥ ⎝ q2 ⎠ ⎢⎣ 2 ⎝ q2 ⎠ ⎦

(13.12)

where A = (R-1)/R. For example, in the cultural grasslands data matrix (see Table 13.3), there were 26 ant species observed in 12 plots, with six species each occurring in exactly one plot, and 8 species each occurring in exactly two plots. Thus, Sobs = 26, q1 = 6, q2 = 8, and R = 12, so Chao2 = 28.45. Sampling additional plots of cultural grasslands should yield a minimum of two or three additional species. The 95% confidence interval (calculated from the variance in Equation 13.12)

23 See Colwell (2011), Appendix B of the EstimateS User’s Guide for other cases and for the calculation of asymmetrical confidence intervals, which are also used in Figure 14.3.

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is from 23.2 to 33.7 species. Table 13.4 summarizes the calculations for the individual-based spider data in each of the four treatments of the hemlock experiment, and Table 13.5 summarizes the calculations for the sample-based Massachusetts ant data in each of the three habitats. The minimum estimated species richness and its confidence interval vary widely among samples. In general, the greater the difference between the observed species richness (Sobs) and the asymptotic estimator (Chao1 or Chao2), the larger the uncertainty and the greater the size of the resulting confidence interval. The most extreme example is from the Hardwood Control treatment in the spider data set. Although 28 species were recorded in a sample of 250 individuals, there were 18 singletons and only one doubleton. The resulting Chao1 = 190 species, with a (parametric) confidence interval from –162 to 542 species! The large number of singletons in these data signals a large number of undetected species, but the sample size is not sufficient to generate an extrapolated estimate with any reasonable degree of certainty. How many additional individuals (or samples) would need to be collected in order to achieve these asymptotic estimators? Equation 13.7 provides a natural “stopping rule”: sampling can stop when there are no singletons in the data set, so that all species are represented by at least two individuals. That amount of sampling often turns out to be very extensive: by the time enough individuals have been sampled to find a second representative for each singleton species, new singletons will have turned up in the data set.

TABLE 13.4 Asymptotic estimator summary statistics for individual-based sampling of spiders Treatment

Sobs

f2 Chao1

2 σChao1

Hemlock Control

106

29.8

Girdled

168

44.0

207

Hardwood Control

250

190.0

32,238

Logged

252

61.5

34.6

346.1

Confidence n* interval (g = 1.0)

n* (g = 0.90)

(18, 41)

345

(6, 72)

1357

355

17,676

4822

2528

609

(–162, 542) (25, 98)

n = number of individuals collected in each treatment; Sobs = observed number of species; f1= number of singleton species; f2 = number of doubleton species; Chao1 = estimated asymptotic species richness; σ 2Chao1= variance of Chao1; confidence interval = parametric 95% confidence interval; n* (g = 1.0) = estimated number of additional individuals that would need to be sampled to reach Chao1; n* (g = 0.90) = estimated number of additional individuals that would need to be sampled to reach 90% of Chao1.

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TABLE 13.5 Asymptotic estimator summary statistics for sample-based collections of ants Habitat

Sobs

Chao2

2 σChao2

Confidence interval

R* (g = 1.0)

R* (g = 0.90)

Cultural grasslands

28.06

5.43

(23, 33)

Oak-hickorywhite pine forest

28.05

5.36

(24, 33)

Successional shrubland

26.1

37.12

(12, 40)

R = number of sampled plots in each habitat; Sobs = observed number of species; q1= number of unique species; q2 = number of duplicate species; Chao2 = estimated asymptotic species richness; σ 2Chao1 = variance of Chao2; confidence interval = parametric 95% confidence interval; R* (g = 1.0) = estimated number of additional samples that would need to be sampled to reach Chao2; R* (g = 0.90) = estimated number of additional samples that would need to be sampled to reach 90% of Chao2.

But what is the number of individuals (or samples) that would be needed to eliminate all of the singletons (or uniques)? Chao et al. (2009) derive formulas (and provide an Excel-sheet calculator) for estimating the number of additional individuals (n*) or samples (R*) that would need to be collected in order to achieve Chao1 or Chao2. These estimates are provided in the last two columns of Tables 13.4 and 13.5. The column for n* when g = 1 gives the sample size needed to achieve the asymptotic species richness estimator, and the column for n* when g = 0.9 gives the sample size needed to achieve 90% of the asymptotic species richness. The additional sampling effort varies widely among the different samples, depending on how close Sobs is to the asymptotic estimator. For example, in the oak-hickory-white pine forest, 26 ant species were collected in 11 sample plots. The asymptotic estimator (Chao2) is 28.1 species, and the estimated number of additional sample plots needed is 17, a 54% increase over the original sampling effort. At the other extreme, 28 spider species were recorded from a collection of 250 individuals in the Hardwood Control treatment of the hemlock experiment. To achieve the estimated 190 species (Chao1) at the asymptote, an additional 17,676 spiders would need to be sampled, a 70-fold increase over the original sampling effort. Other biodiversity samples typically require from three to ten times the original sampling effort in order to reach the estimated asymptotic species richness (Chao et al. 2009). The effort is usually considerable because a great deal of sampling in the thin right-hand tail of the species-abun-

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dance distribution is necessary to capture the undetected rare species. The sampling requirements are less burdensome if we can be satisfied with capturing some lesser fraction, say 90%, of the asymptotic species richness. For example, in the Hemlock Control treatment, 23 spider species were observed, with an estimated asymptote of 29.8. An additional 345 individual spiders would need to be censused to reach this asymptote, but if we would be satisfied with capturing 90% of asymptotic richness, only an additional 65 spiders would be needed. Rarefaction Curves Redux: Extrapolation and Interpolation

Biodiversity sampling begins with a reference sample—a standardized collection of individuals (or samples) that has a measured number of species. With rarefaction, the data are interpolated to progressively smaller sample sizes to estimate the expected species richness. With asymptotic estimators, the same data are extrapolated to a minimum asymptotic estimator of species richness, with an associated sampling effort that would be needed to achieve that level of richness. Recently, Colwell et al. (2012) unified the theoretical frameworks of rarefaction and asymptotic richness estimation. They derived equations that link the interpolated part of the rarefaction curve with the extrapolated region out to the asymptotic estimator. In standard rarefaction, the variance is conditional on the observed data, and the confidence interval converges to zero at the observed sample size (see Figures 13.2 and 13.4). In the Colwell et al. (2012) framework, the rarefaction variance is unconditional. The reference sample is viewed more properly as a sample from a larger assemblage, and the unconditional variance can be derived based on the expectation and variance of the asymptotic estimator. We will forego the equations here, but Figure 13.6 illustrates these extended rarefaction/extrapolation curves for the Massachusetts ant data. These curves graphically confirm the results of the rarefaction analysis (see Figure 13.4), which is that ant species richness is fairly similar in all three habitats. However, the extrapolation of the successional shrublands data out to an asymptotic estimator is highly uncertain and generates a very broad confidence interval because it was based on a reference sample of only five plots.

Estimating Species Diversity and Evenness Up until now, this chapter has addressed the estimation of species richness, which is of primary concern in many applied and theoretical questions. Focusing on species richness seems to ignore differences in the relative abundance of species, although both the asymptotic estimators and the shape of the rarefaction curve depend very much on the commonness versus rarity of species.

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Estimating Species Diversity and Evenness

Figure 13.6 Interpolated and extrapo-

lated sample-based rarefaction curves of the ant data. Light gray: oak-hickorywhite pine forests; dark gray = cultural grasslands; blue = successional shrublands. The filled points are the actual samples and the open points are the extrapolated species richness (Chao2). The solid lines are the interpolated regions of each curve and the dashed lines are the extrapolated regions of each curve. The shaded regions indicate the approximate 95% confidence interval for each curve.

30 Species richness

477

15 20 Number of samples

Ecologists have tried to expand the measure of species diversity to include components of both species richness and species evenness. Consider two forests that each consist of five species and 100 individual trees. In Forest A, the 100 trees are apportioned equally among the five species (maximum evenness), whereas, in Forest B, the first species is represented by 96 trees, and the remaining four species are represented by one tree each. Most researchers would say that Forest A is more diverse than Forest B, even though both have the same number of species and the same number of individuals. If you were to walk through Forest A, you would frequently encounter all five species, whereas, in Forest B, you would mostly encounter the first species, and rarely encounter the other four species.24

24 The great explorer and co-discoverer of evolution, Alfred Russel Wallace (1823–1913) was especially impressed by the high diversity and extreme rarity of tree species in Old World tropical rainforests: “If the traveler notices a particular species and wishes to find more like it, he may often turn his eyes in vain in every direction. Trees of varied forms, dimensions and colours are around him, but he rarely sees any one of them repeated. Time after time he goes towards a tree which looks like the one he seeks, but a closer examination proves it Alfred Russel to be distinct. He may at length, perhaps, meet with a second speciWallace men half a mile off, or may fail altogether, till on another occasion he stumbles on one by accident” (Wallace 1878). Wallace’s description is a nice summary of the aspect of diversity that is measured by PIE, the probability of an interspecific encounter (see Equation 13.15; Hurlbert 1971).

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Many diversity indices try to incorporate effects of both species richness and species evenness. The most famous diversity index is the Shannon diversity index, calculated as: s

H ′ = − ∑ pi log ( pi )

(13.13)

i =1

where pi is the proportion of the complete assemblage represented by species i(pi=Ni /N). The more species in the assemblage and the more even their relative abundances, the larger H ′will be. There are literally dozens of such indices based on algebraic transformations and summations of pi. In most cases, these diversity indices do not have units that are easy to interpret. Like species richness, they are sensitive to the number of individuals and samples collected, and they do not always have good statistical performance. An exception is Simpson’s (1949) index of concentration: s

D = ∑ pi2

(13.14)

i =1

This index measures the probability that two randomly chosen individuals represent the same species. The lower this index is, the greater the diversity. Rearranging and including an adjustment for small sample sizes, we have: PIE =

s ⎞ n ⎛ 1.0 − pi2 ⎟ ∑ ⎜ (n − 1) ⎝ ⎠ i =1

(13.15)

PIE is the probability of an interspecific encounter (Hurlbert 1971), the probability that two randomly chosen individuals from an assemblage will represent two different species.25 There are 3 advantages of using PIE as a simple diversity 25

In economics, the PIE index (without the correction factor n/(n-1)) is known as the Gini coefficient (see Equation 11.26; Morgan 1962). Imagine a graph in which the cumulative proportions of species (or incomes) are plotted on the y-axis, and the rank order of the species (in increasing order) is plotted on the x-axis. If the distribution is perfectly even, the graph will form a straight line. But if there is any deviation from perfect evenness, a concave curve, known as the Lorenz (1905) curve, is formed with the same starting and ending point as the straight line. The Gini coefficient quantifies inequality in income as the relative area between the straight line and the Lorenz curve. Of course, for income distributions, the absolute difference between the richest and the poorest classes may be more important than the relative evenness of the different classes. Ecologists have used the Gini coefficient to quantify the magnitude of competitive interactions in dense populations of plants (Weiner and Solbrig 1984).

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Estimating Species Diversity and Evenness

index. First, it has easily interpretable units of probability and corresponds intuitively to a diversity measure that is based on the encounter of novel species while sampling (see Footnote 24 in this chapter). Second, unlike species richness, PIE is insensitive to sample size; a rarefaction curve of the PIE index (using the estimator pˆ i =Xi /n generates a flat line. Finally, PIE measures the slope of the individual-based rarefaction curve measured at its base (Olsweski 2004). Returning to the example of two hypothetical forests, each with five species and 100 individuals, the maximally even Forest A (20, 20, 20, 20, 20) has PIE = 0.81, whereas the maximally uneven Forest B (96, 1, 1, 1, 1) has PIE = 0.08. In spite of the advantages of the PIE index as an intuitive measure of species diversity based on relative abundance, the index is bounded between 0 and 1, so that the index becomes “compressed” near its extremes and does not obey a basic doubling property: if two assemblages with the same relative abundance distribution but no species in common are combined with equal weight, diversity should double. Certainly species richness obeys this doubling property. However, if we double up Forest A (20, 20, 20, 20, 20) to form Forest AA (20, 20, 20, 20, 20, 20, 20, 20, 20, 20), diversity changes from PIEA = 0.81 to PIEAA = 0.90. If we double up Forest B (96, 1, 1, 1, 1) to form Forest BB (96, 96, 1, 1, 1, 1, 1, 1, 1, 1), diversity changes from PIEB = 0.08 to PIEBB = 0.54. Hill Numbers

Hill numbers (Hill 1973a) are a family of diversity indices that overcome the problems of many of the diversity indices most commonly used by ecologists. Hill numbers preserve the doubling property, they quantify diversity in units of modified species counts, and they are equivalent to algebraic transformations of most other indices. Hill numbers were first proposed as diversity measures by the ecologist Robert MacArthur (MacArthur 1965; see Footnote 6 in Chapter 4), but their use did not gain much traction until 40 years later, when they were reintroduced to ecologists and evolutionary biologists in a series of important papers by Jost (2006, 2007, 2010). The general formula for the calculation of a Hill number is: q

(

D = ∑ is=1 piq

)(

1/ 1− q )

(13.16)

As before, pi is the “true” relative frequency of each of each species (pi = Ni / N, for i = 1 to S) in the complete assemblage. The exponent q is a non-negative integer that defines the particular Hill number. Changing the exponent q yields a family of diversity indices. As q increases, the index puts increasing weight on the most common species in the assemblage, with rare species making less and less of a contribution to the summation. Once q ⲏ 5 , Hill numbers rapidly converge to the inverse of the relative abundance of the most common species. Negative values for

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q are theoretically possible, but they are never used as diversity indices because they place too much weight on the frequencies of rare species, which are dominated by sampling noise. As q increases, the diversity index decreases, unless all species are equally abundant (maximum evenness). In this special case, the Hill number is the same for all values of q, and is equivalent to simple species richness. Regardless of the exponent q, the resulting Hill numbers are always expressed in units of effective numbers of species: the equivalent number of equally abundant species. For example, if the observed species richness in a sample is ten, but the effective number of species is five, the diversity is equivalent to that of a hypothetical assemblage with five equally abundant species.26 The first three exponents in the family of Hill numbers are especially important. When q = 0, 1

(

⎛ s ⎞ D = ⎜ ∑ pi0 ⎟ = S ⎝ i =1 ⎠

(13.17)

)

because pi0 = 1 , 0 D = ∑ i =11 , and S1 = S. Thus, 0D corresponds to ordinary species richness. Because 0D is not affected by species frequencies, it actually puts more weight on rare species than any of the other Hill numbers. S

For q = 1, Equation 13.16 cannot be solved directly (because the exponent of the summation, 1/(1 – q) is undefined), but in the limit it approaches: 1

D=e

H′

⎛− ⎜⎝

∑ i =1 pi log p i ⎞⎟⎠ s

(13.18)

Thus 1D is equivalent to the exponential of the familiar Shannon measure of diversity (H ′) (see Equation 13.13). 1D weights each species by its relative frequency. Finally, for q = 2, Equation 13.16 gives: 2

⎛ s ⎞ D = ⎜ pi2 ⎟ ⎝ i=1 ⎠

∑

−1

(∑

1 s p2 i=1 i

)

(13.19)

The effective number of species is very similar in concept to the effective population size in population genetics, which is the equivalent size of a population with entirely random mating, with no diminution caused by factors such as bottlenecks or unequal sex ratios (see Footnote 3 in Chapter 3). The concept of effective number of species also has analogues in physics and economics (Jost 2006).

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Software for Estimation of Species Diversity

Thus 2D is equivalent to the inverse of Simpson’s index (see Equation 13.14). 2D and qD for values of q > 2 give heavier weight to the more common species. Hill numbers provide a useful family of diversity indices that consistently incorporate relative abundances while at the same time express diversity in units of effective numbers of species. Two caveats are important when using Hill numbers, however. First, no diversity index can completely disentangle species richness from species evenness (Jost 2010). The two concepts are intimately related: the shape of the rarefaction curve is affected by the relative abundance of species, and any index of evenness is affected by the number of species present in the assemblage. Second, Hill numbers are not immune to sampling effects. Our treatment of Hill numbers is based on the true parameters (pi and S) from the complete assemblage, although calculations of Hill numbers are routinely based on estimates of those parameters ( pˆ i and Sobs) from a reference sample.27 Species richness is the Hill number with q = 0, and this chapter has emphasized that both the number of individuals and the number of samples have strong influences on Sobs. Sample size effects are important for all the other Hill numbers, although their effect diminishes as q is increased. Figure 13.7 depicts rarefaction curves for the spider data of the Hardwood Control, and illustrates the sampling effect with the Hill numbers 0D, 1D, 2D, and 3D.

Software for Estimation of Species Diversity Software for calculating rarefaction curves and extrapolation curves for species richness and other diversity indices is available in Anne Chao’s program SPADE (Species Prediction and Diversity Estimation: chao.stat.nthu.edu.tw/softwareCE.html), and in Rob Colwell’s program EstimateS (purl.oclc.org/estimates). The vegetarian library in R, written by Noah Charney and Sydne Record, contains many useful functions for calculating Hill numbers (cran.r-project.org/ web/packages/vegetarian/index.html). R code used to analyze and graph the data in this chapter is available in the data section of this book’s website (harvardforest.fas.harvard.edu/ellison/publications/primer/datafiles) and as part of

27 The problem with using the standard “plug-in” estimator pˆ i = X i / n is that, on average, it over-estimates pi for the species that are present in the reference sample. Chao and Shen (2003) derive a low-bias estimator for the Shannon diversity index (see Equation 13.13) that adjusts for the both biases in pˆ i and for the missing species in the reference sample. Anne Chao and her colleagues are currently developing asymptotic estimators and variances for other Hill numbers.

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CHAPTER 13 The Measurement of Biodiversity

Figure 13.7 Individual-based rarefaction

30 q=0 25 Effective number of species

curves for the family of Hill numbers (see Equation 13.16) with exponents q = 0 to 3. Each curve is based on 1000 randomization of the Hardwood Control reference sample of 250 spiders and 28 species from the Harvard Forest Hemlock Removal Experiment (see Table 13.1). Note that q = 0 is simple species richness, so the curve is identical to the light gray rarefaction curve shown in Figure 13.3.

20 15 10 q=1 q=2 q=3

150 200 100 Number of individuals

250

300

EcoSimR, a set of R functions and scripts for null model analysis (www.uvm.edu/ ~ngotelli/EcoSim/EcoSim.html).

Summary The measurement of biodiversity is a central activity in ecology. However, biodiversity data (samples of individuals, identified to species, or records of species incidences) are labor-intensive to collect, and in addition usually represent only a tiny fraction of the actual biodiversity present. Species richness and most other diversity measures are highly sensitive to sampling effects, and undetected species are a common problem. Comparisons of diversity among treatments or habitats must properly control for sampling effects. Rarefaction is an effective method for interpolating diversity data to a common sampling effort, so as to facilitate comparison. Extrapolation to asymptotic estimators of species richness is also possible, although the statistical uncertainty associated with these extrapolations is often large, and enormous sample sizes may be needed to achieve asymptotic richness in real situations. Hill numbers provide the most useful diversity measures that incorporate relative abundance differences, but they too are sensitive to sampling effects. Once species diversity has been properly estimated and sampling effects controlled for, the resulting estimators can be used as response or predictor variables in many kinds of statistical analyses.

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CHAPTER 14

Detecting Populations and Estimating their Size

We began this book with an introduction to probability and sampling and the central tasks of measurement and quantification. Questions such as “Are two populations different in size?”, “What is the density of Myrmica ants in the forest?”, or “Do these two sites have different numbers of species?” are fundamental to basic ecological studies and many applied problems in natural resource management and conservation. The different inferential approaches (see Chapter 5) and methods for statistical estimation and hypothesis testing (see Chapters 9–13) that we have discussed up until now uniformly assume not only that the samples are randomized, replicated, and independent (see Chapter 6), but also that the investigator has complete knowledge of the sample. In other words, there are no errors or uncertainty in the measurements, and the resulting data actually quantify the parameters in which we are interested. Are these reasonable assumptions? Consider the seemingly simple problems of comparing the nest density (see Chapter 5) or species richness (see Chapter 13) of ground-foraging ants in fields and forests. The measurements for these examples include habitat descriptors and counts of the number of ant nests per sampled quadrat (see Table 5.1) or the number of species encountered in different sites (see Table 13.5). We used these data to make inferences about the size of the population or the species richness of the assemblage, which are the parameters we care about. We can be confident of our identifications of habitats as forests or fields, but are we really certain that there were only six ant nests in the second forest quadrat or 26 species in the average oak-hickory-white pine forest? What about that Stenamma brevicorne nesting in between two decomposing leaves that we missed, or the nocturnal Camponotus castaneus that was out foraging while we were home sleeping? How easy would it be to count all of the individuals accurately, or even to find all of the species that are present?

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CHAPTER 14 Detecting Populations and Estimating their Size

This chapter introduces methods to address the problem that the probability of detecting all of the individuals in a sample or all of the species in a habitat is always less than 1. Because sampling is imperfect and incomplete, the observed count C is always less than the true number of individuals (or species) N. If we assume that there is a number p equal to the fraction of individuals (or species) of the total population that we sampled, then we can relate the count C to the total number N as: C = Np

(14.1)

If we had some way to estimate p, we could then estimate the total population size (or number of species) N, given our observed count C: Nˆ = C / pˆ

(14.2)

As in earlier chapters, the “hats” (ˆ) atop N and p in Equation 14.2 indicate that these values are statistical estimates of the unknown, true values of these parameters. As we develop models in this chapter based on Equations 14.1 and 14.2, we will usually add one or more subscripts to each of C, N, and p to indicate a particular sample location or sample time. Isolating Nˆ on the left-hand side of Equation 14.2 indicates that we are explicitly interested in estimating the total number of objects (see Chapter 13)—e.g., individuals in a population, species in an assemblage, etc. But Nˆ is not the only unknown quantity in Equation 14.2. We first need to estimate pˆ , the proportion of the total number of objects that were sampled. In Chapter 13, we faced a similar problem in estimating the proportional abundance of a given species as pˆi = Xi /n, where Xi was the number of individuals of species i and n was the number of individuals in the reference sample, which was always less than the total number of individuals in the assemblage (N). The rearrangement of Equation 14.2 highlights the necessity of estimating both pˆ and Nˆ . In summary, a simple idea—that we are unable to have seen (or counted) everything we intended to see or count (as in Figure 14.1)—and its encapsulation in a simple equation (Equation 14.2), brings together many of the ideas of probability, sampling, estimation, and testing that we have presented in the preceding chapters. The apparent simplicity of Equation 14.2 belies a complexity of issues and statistical methods. We begin by estimating the probability that a species is present or absent at a site (occupancy), and then move on to estimate

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Occupancy

485

Figure 14.1 Sample replication and detection probability. Within a spatially defined sample space (large circle), only a subset of plots can be sampled (blue squares). Within each plot, individuals of interest (circles) will either be found (detected, indicated by black circles) or not found (undetected, indicated by white circles). (After MacKenzie et al. 2006.)

how many individuals of that species occur at the site (abundance), given that (i.e., conditional on) it is present.1

Occupancy Before we can estimate the size of a local population, we need to determine whether the species is present at all; we refer to a site where a species is present as an occupied site, and one where a species is absent as an unoccupied site. We define occupancy as the probability that a sampling unit or an area of interest (henceforth, a site) that is selected at random contains at least one individual of the species of interest. In brief, occupancy is the probability that a species is present at a site. If we sample x sites out of a total of s possible sites,2 we can estimate occupancy as: 1 Although it is logical to first ask whether a species is present or not, and then ask how many individuals of a given species there are, methods for estimating the latter were actually developed first. We suspect that estimates of population size were derived before estimates of occupancy because mark-recapture methods were initially applied to common species of commercially important animals, such as fish or deer. There was no question these animals were present, but their population sizes needed to be estimated. Later, biologists became interested in occupancy models as a tool for studying species that are too rare to find and count easily. Unfortunately, many of the species that are currently extinct or very rare were formerly common, but have been overexploited by hunting or harvesting. 2

Be aware that the notation among the chapters of this book is not consistent. In the species richness literature (see Chapter 13), s refers to species, and n refers to sites or samples. But in the occupancy literature, s refers to samples, and n refers to species or individuals. In both of these chapters, we have stuck with the variable notation that is most commonly used in cited papers in these subdisciplines rather than imposing our own scheme to keep the notation consistent among chapters. Caveat lector !

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CHAPTER 14 Detecting Populations and Estimating their Size

x ψˆ = s

(14.3)

If there is no uncertainty in detection probability (the value of pˆ in Equation 14.2), then we have a simple estimation problem. We start by assuming that there are no differences in occupancy among all of the sampled sites (that is ψ1 = ψ2 = … = ψs ≡ ψ) and that we can always detect a species if it is indeed present (the detection probability p = 1). In this simple case, the presence or absence of an individual species at a sampled site is a Bernoulli random variable (it is either there or it is not; see Chapter 2), and the number of sites in which the species is present, x, will be a binomial random variable with expected value given by Equation 14.3 and variance equal to ψ(1 – ψ)/s. If the detection probability p < 1, but is still a known, fixed quantity, then the probability of detecting the species in at least one of t repeated surveys is p′ = 1− (1 − p ) × (1 − p ) × … × (1 − p ) = 1 − (1 − p )

(14.4)

t times

Equation 14.4 is simply 1 minus the probability of not having detected the species in all t of the surveys. With the detection probability p′ now adjusted for multiple surveys, we can move on to estimating occupancy, again as a binomial random variable. Equation 14.3 is modified to adjust occupancy by the detection probability given by Equation 14.4, resulting in: x ψˆ = sp′

(14.5)

Note that the numerator x in Equation 14.5 is the number of sites in which the species is detected, which is less than or equal to the number of sites that the species actually occupies. The variance of this estimate of occupancy is larger than the variance of the estimate of occupancy when the detection probability equals one: Var ( ψˆ ) =

1 − ψp′ ψ (1 − ψ ) ψ (1 − p′ ) = + sp′ s sp′

(14.6)

The first term on the right-hand side of Equation 14.6 is the variance when p = 1, and the second term is the inflation in the variance caused by imperfect detection. The real challenge, however, arises when detection probability p is unknown, so now the number of sampled sites in which the species is actually present also has to be estimated: xˆ ψˆ = s

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(14.7)

Occupancy

Because there is uncertainty in both detection probability and occupancy, 3 Equation 14.7 is the most general formula for estimating occupancy. Two approaches to this have been developed to estimating occupancy and detection probabilities. In one approach, we first estimate detection probability and then use Equation 14.5 to estimate occupancy. In the second approach, both occupancy and detection are estimated simultaneously with either a maximum likelihood or a Bayesian model (see Chapter 5). Although the first approach is computationally easier (and usually can be done with pencil and paper), it assumes that occupancy and detection probabilities are simple constants that do not vary in time or space. The second approach can accommodate covariates (such as habitat type or time of day) that are likely to affect occupancy or detection probabilities. With the second approach, we can also compare the relative fit of models for the same data set that incorporate different covariates. Both approaches are described in detail by MacKenzie et al. (2006); we present only the second approach here. The Basic Model: One Species, One Season, Two Samples at a Range of Sites

The basic model for jointly estimating detection probabilities and occupancy was developed by MacKenzie et al. (2002). In our description of occupancy models, we continue to use the notation introduced in Equations 14.3–14.7: occupancy (ψ) is the probability that a species is present at site i; pit is the probability that the species is detected at site i at sample time t; T is the total number of sampling times; s is the total number of sites surveyed; xt is the number of sites

For charismatic megafauna such as pandas, mountain lions, and Ivory-billed Woodpeckers that are the focus of high-profile hunting, conservation, or recovery efforts, occupancy probabilities may be effectively 0.0, but uncertainty in detection is very high and unconfirmed sightings are unfortunately the norm. The Eastern Mountain Lion (Puma concolor couguar) was placed on the U.S. Fish & Wildlife Service’s Endangered Species List (www.fws.gov/endangered/) in 1973 and is now considered extinct; the last known individual was captured in 1938 (McCollough 2011). Similarly, the last specimen of the Ivory-billed Woodpecker (Campephilus principalis) was collected in the southeastern United States in 1932, and the last verified sighting was in 1944. But both species are regularly “rediscovered.” For the Ivory-billed Woodpecker (Gotelli et al. 2012), the probability of these rediscoveries being true positives (< 6 × 10–5) is much smaller than the probability that physicists have falsely discovered the Higgs boson after 50 years of searching. Hope springs eternal, however, and signs and sightings of both Eastern Mountain Lions in New England and Ivory-billed Woodpeckers in the southeastern U.S. are regularly reported—but never with any verifiable physical or forensic evidence (McKelvey et al. 2003). There is also even a fraction of the tourist economy that caters to the searchers. Bigfoot in the Pacific Northwest and “The King” in Graceland now have lots of company!

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Chapter 14 Detecting Populations and Estimating their Size

for which the species was detected at time t; and x is the total number of sites at which the species was detected on at least one of the sampling dates. The simplest model for estimating occupancy requires at least two samples from each of one or more sites. In addition, we assume that, for the duration of all of our samples, the population is closed to immigration, emigration, and death: that is, individuals neither enter nor leave a site between samples (we relax this closure assumption in a subsequent section). We also assume that we can identify accurately the species of interest; false positives are not allowed. However, false negatives are possible, with probability equal to [1 – occupancy]. Finally, we assume that all sites are independent of one another, so that the probability of detecting the species at one site is not affected by detections at other sites. In Chapter 9, we illustrated how the number of New England forest ant species changes as a function of latitude and elevation. In addition to sampling ants in forests, we also sampled ants in adjacent bogs, which harbor a number of ant species specialized for living in sunny, water-logged habitats. In the summer of 1999, we sampled each of 22 bogs twice, with samples separated by approximately six weeks. The data for a single species, Dolichoderus pustulatus, which builds carton nests inside of old pitcher-plant leaves, are shown in Table 14.1 (from Gotelli and Ellison 2002a). We collected D. pustulatus at x = 16 of the s = 22 bogs (x1 = 11 detections in the first sample, x2 = 13 detections in the second sample). We are interested in estimating ψ, the probability that a randomly chosen bog was indeed occupied by D. pustulatus. For each site at one of the two sampling dates, D. pustulatus was either detected (1) or not (0). If it was detected on only one of the two dates, we assumed it was present but not detected at the other date (because of the closure assumption of this model). The detection history for a single site can be written as a 2-element string of ones and zeros, in which a 1 indicates that the individual was detected (collected) and a 0 indicates that it was not. Each site in this example has one of four possible detection histories, each with 2 elements: (1,1)—D. pustulatus was collected in both samples; (1,0)—D. pustulatus was collected only in the first sample; (0,1)—D. pustulatus was collected only in the second sample; and (0,0)—D. pustulatus was collected in neither sample. Figure 14.2 illustrates the sampling histories for all 22 sites. If we collected D. pustulatus at site i in the first sample, but not in the second (sampling history [1,0]), the probability that we would detect it at that site is: P ( xi ) = ψ i pi1 (1 − pi 2 )

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(14.8)

Occupancy

489

Ants present Ants absent

Bog site

Figure 14.2 The collection histories of the ant species Dolichoderus pustulatus at 22 bogs (y-axis) on each of two sampling dates (x-axis). Blue indicates this species was collected and magenta that it was not (data from columns 2 and 3 of Table 14.1). If we assume that the populations of ants at each site were closed, detection was imperfect, because at eight of the 22 sites the species was detected on one sampling date but not the other. At eight sites it was detected on both sampling dates, and at the remaining six sites it was detected on neither sampling date.

First observation

Second observation

In words, this probability is equal to D. pustulatus occupying the site (ψi) and being detected at time 1 (pi1) and not being detected at time 2 (1 – pi2). On the other hand, if we never collected D. pustulatus at site j, the collection history would be (0,0), and the probability that we did not detect it would be the sum of the probabilities that it did not occupy the site (1 – ψj), or it did occupy the site (ψj) and was not collected at either time (∏ t2=1(1 − p jt )) :

( ) (

)

(

P x j = 1 − ψ j + ψ j ∏ t2=1 1 − p jt

)

(14.9)

In general, if we assume that both occupancy and detection probabilities are the same at every site, we can combine the general forms of Equations 14.8 and 14.9 to give the full likelihood: T T ⎡ ⎤ ⎡ ⎤ L ( ψ , p1 , … , px ) = ⎢ ψ x ∏ ptxt (1 − pt )x − xt ⎥ × ⎢(1 − ψ) + ψ ∏ (1 − pt ) ⎥ t =1 ⎣ t =1 ⎦ ⎣ ⎦

s− x

(14.10)

For the data in Table 14.1, s = 22, x1 = 11, x2 = 13, x = 16, and T = 2. In this context, the likelihood is the value of ψ that would be most probable (= likely), given the data on detection histories (p1, … , px).

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CHAPTER 14 Detecting Populations and Estimating their Size

TABLE 14.1 Occurrence of Dolichoderus pustulatus at 22 bogs in Massachusetts and Vermont Site

Sample1 Sample2 Latitude Elevation

Bog area Date1 Date2

ARC

42.31

1190

153

195

42.56

274

105369

160

202

CAR

44.95

133

38023

153

295

42.05

210

73120

181

216

CHI

44.33

362

38081

174

216

CKB

42.03

152

7422

188

223

COL

44.55

623284

160

202

HAW

42.58

543

36813

175

217

HBC

42.00

11760

191

241

MOL

44.50

236

8852

153

195

MOO

44.76

353

864970

174

216

42.23

491

89208

174

209

PEA

44.29

468

576732

160

202

PKB

42.19

491189

188

223

42.57

335

40447

160

202

42.17

10511

174

209

SKP

42.05

55152

191

241

SNA

44.06

313

248

167

209

SPR

43.33

158

435

167

209

SWR

42.27

121

19699

153

195

TPB

41.98

389

2877

181

216

WIN

42.69

323

84235

167

202

Total occurrences

Values shown are the name of each bog (site); whether D. pustulatus was present (1) or absent (0) on the first (sample1) or second (sample2) sample date; the latitude (decimal degrees), elevation (meters above sea level) and area (m2) of the bog mat; and the days of the year (January 1 1999 = 1) on which each bog was sampled (date1, date2).

In addition, we can model both occupancy and detection probability as a function of different covariates. Occupancy could be a function of site-based covariates such as latitude, elevation, bog area, or average annual temperature. In contrast, detection probability is more likely to be a function of time-based

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Occupancy

covariates that are associated with each census. Such time-based covariates might include the day of the year on which a site was sampled, the time of day we arrived at the site, or weather conditions during sampling. In general, both occupancy and detection probability can be modeled as functions of either site-based or time-based covariates. Once the model structure is selected, the covariates can be incorporated into the model using logistic regression (see Chapter 9): y=

exp (b X site )

(14.11a)

exp (b X time )

(14.11b)

1 + exp (b X site )

1 + exp (b X time )

In these two equations, the parameters and covariates are bold-faced because they represent vectors, and the regression model can be specified using matrix operations (see Appendix Equations A.13 and A.14). If we include covariates in our estimation of occupancy, then we estimate the average occupancy as: s ∑i =1 yˆ i ˆ y= s

(14.12)

We used Equations 14.10–14.12 to estimate the occupancy of Dolichoderus in Massachusetts and Vermont bogs.4 We first estimated occupancy without assuming the influence of any covariates, then fit three alternative models with different sets of covariates: (1) a model in which detection probability varied with time of sample (sampling date); (2) a model in which occupancy varied with three site-specific covariates (latitude, elevation, and area); and (3) a model in which occupancy varied with the three site-specific geographic covariates and detection probability varied with time of sample. We compared the fit of the four different models to the data using Akaike’s Information Criteria (AIC; see “Model Selection Criteria” in Chapter 9). The results, shown in Table 14.2, suggest that the best model is the simplest one—constant detection probability at the two sample dates and occupancy that does not co-vary geographically—but that the model that includes a temporally varying detection probability fits almost as well (the difference in the AIC 4 All of these models were fit using the occu function in the R library unmarked (Fiske and Chandler 2011). Spatial and temporal covariates were first rescaled as Z scores (see Chapter 12).

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TABLE 14.2 Estimates of occupancy and detection probability of Dolichoderus pustulatus in 22 New England bogs Model 1 Description

Number of parameters to estimate

No covariates

Model 2

Model 3

Model 4

Detection Occupancy Detection varies probability varies with bog with time and varies with geographic occupancy time of sample characteristics varies geographically 3

Estimated occupancy (ψˆ )

0.82 (0.13)

0.83 (0.13)

0.80 (0.10)

0.88 (0.11)

Estimated detection probability (pˆ)

0.67 (0.11)

0.66 (0.12)

0.76 (0.09)

0.67 (0.10)

AIC

63.05

64.53

66.45

68.03

Dark blue shading indicates the best-fitting models as determined by AIC; numbers in parentheses are the standard error of the given parameter.

between these two models is less than 2). All four of the models estimated occupancy at between 80% and 90%, which suggests that we missed finding D. pustulatus in at least two of the bogs that were sampled. Note that even the simplest model without any covariates requires that we estimate two parameters—occupancy and detection (see Equation 14.7). Occupancy is of primary interest, whereas detection probability is considered a nuisance parameter—a parameter that is not of central concern, but that we nonetheless must estimate to get at what we really want to know. Our estimates of detection probability ranged from 67% in the simple models to 76% in the model in which occupancy was a function of three geographic covariates. Even though we have been studying pitcher plants and their prey, and collecting ants in bogs for nearly 15 years, we can still overlook common ants that build their nests inside of unique and very apparent plants. The standard errors in Table 14.2 of all of the occupancy estimates of D. pustulatus were relatively large because only two samples (the absolute minimum necessary) were used to fit the model. With a greater number of sampling times, the uncertainty in the estimate of occupancy would decrease. In simulated data sets, for a sample size s of 40 sites with detection probabilities (p) of 0.5, at least five samples are needed to get accurate estimates of occupancy (MacKenzie et

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Occupancy

al. 2002). In our example, the standard error of occupancy could have been reduced by as much as 50% if we had had visited each bog five times instead of two. We return to a discussion of general issues of sampling at the end of this chapter. Occupancy of More than One Species

....

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In Chapter 13, we described asymptotic species richness estimators (Chao1 and Chao2) which are used to estimate the number of undetected species in a reference sample. These estimators are based on the numbers of singletons, or uniques, and doubletons, or duplicates (species in the reference sample represented by exactly one or two individuals, respectively; see Equations 13.7 and 13.8). We also estimated the number of additional samples that would be needed to find all of the undetected species (see Tables 13.4 and 13.5; Chao et al. 2009). As with rarefaction, these estimators assume that the probability of detection per individual is constant and does not differ among individuals of different species. Therefore, the detection probability of different species is proportional to their relative frequency in the assemblage (see Equations 13.1–13.4). However, it would be more realistic to assume that occupancy and detection probabilities vary among species, through time, and as a function of site-based variables, and to incorporate these factors into species richness estimators (or other diversity measures). The simple occupancy model for a single species (see Equations 14.7–14.12) can be expanded to incorporate multiple species and used to estimate the number of undetected species. Our ant data from New England bogs, of which the D. pustulatus example (see Tables 14.1 and 14.2) are a subset, provide an illustrative example (Dorazio et al. 2011). At each of the 22 sites in Table 14.1, on each of the two sampling dates, we collected ant occurrence data from a grid of 25 pitfall traps. For the s = 22 sites, the TABLE 14.3 Observed collection occurrence data can be summarized as an n (species) × data of ants in New England bogs s (sites) matrix of observations yik, where i = {1, …, n}, Species i Abundance at site k k = {1, …, s}, and yik is the number of pitfall traps in which each ant species was captured at the kth of the 1 y11 y12 … y1s 22 sites, summed over the two sampling times (Table 2 y21 y22 … y2s 14.3). Thus, yik is an integer between 0 (the species was never collected in a site) and 50 (in both sample periods, the species was found at site k in every one of the 25 pitn yn1 yn2 … yns fall traps in the grid). We observed n species among all the s sites, but we are interested in the total number N Each of n species was observed in yik 25 pitfall traps set at site k (one of 22 bogs). species present at all sites that could have been captured. Because 25 pitfall traps were set at each bog on each of two dates, 0 ≤ yik ≤ 50. N is unknown, but probably includes some species that

CHAPTER 14 Detecting Populations and Estimating their Size

Figure 14.3 Observed (black open circles) and Chao2 estimates (blue filled circles) of species richness of ants in each of 22 New England bogs as a function of their elevation. Vertical lines are 95% confidence intervals (from Equation 13.12). The width of confidence intervals equals zero when doubletons are present in the collection but singletons are not present in the collection.

15 Number of species

494

0 0

100

200 300 400 Elevation (meters)

500

were present but undetected at all of the sites. Although N is at least equal to n, it probably exceeds it (n ≤ N). In this data set, n = 19 species were detected among 22 bogs. Treating captures in pitfall traps as incidence data (Gotelli et al. 2011), the Chao2 estimate (see Equations 13.10–13.12) for regional diversity is 32 species, with a confidence interval of 8.5–54.9. Chao2 estimates of species richness at each bog are shown in Figure 14.3. As discussed in Chapter 13, if there are no uniques in the data set (species recorded at only a single site), the Chao2 estimator equals the observed species richness: there are estimated to be no undetected species, and both the variance and confidence interval equal zero (see Footnote 23 in Chapter 13). However, this index (and others; see Chapter 13) does not incorporate variation among species, sites, or times in occupancy or detection probabilities. Including occupancy, detection probability, and time- and site-specific covariates into species richness estimation is a formidable problem that is just beginning to be addressed (e.g., Dorazio and Royle 2005; MacKenzie et al. 2006; Royle and Dorazio 2008; Dorazio et al. 2011; Dorazio and Rodríguez 2012; Royle and Dorazio 2012). It is important, however, to keep in mind the common goal of both the methods presented in Chapter 13, and the methods presented here: estimation of the total number of N species present at all sites that could be found. Estimation of N is a multi-step process. First, we assume that there are some number of species, N – n, which were present in the bog but that we did not collect. Thus, we can expand the n × s sample matrix (see Table 14.3) to an N × s matrix, in which the unobserved species are designated by additional rows with

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Occupancy

TABLE 14.4 Expanded matrix to include both the observed collection data of ants in New England bogs and the (N – n) species that were present but not observed Species i

Abundance at site k

y11

y12

…

y1s

y21

y22

…

y2s

....

yn1

yn2

…

yns

n+1

…

n+2

…

....

matrix entries assigned to equal zero, as shown in Table 14.4. Second, we assume that the N species in each bog (our alpha diversity; see Chapter 13) are part of a regional species pool of size M (the gamma diversity M >> N), and expand the matrix accordingly to an M × s matrix; see Table 14.5 for this further step. M puts a finite upper bound on regional species richness and constrains subsequent analyses.5 Third, we identify from the regional pool of M species those that can or cannot occur at a site (perhaps due to physiological tolerances, special habitat requirements, etc.) and provide a dummy variable for each species (wi = 1 if it can occur, 0 if it cannot). Finally, we complete the matrix in Table 14.5 with a N × s set of site-specific and species-specific occurrences zik that is equivalent to a species × site incidence matrix in which rows are species, columns are sites, and cell entries are either 1 (present at a site) or 0 (absent at a site) (cf. Table 13.3). In Chapter 13, incidence (or abundance) matrices were fixed observations, but in the framework of occupancy models, the incidence matrix is itself a random variable.

…

The dark blue portion of the matrix is the observed collection data (see Table 14.3), and the light blue portion represents species that could have been present but were not collected.

A Hierarchical Model for Parameter Estimation and Modeling

Putting all of these ideas together yields a type of nested or hierarchical model (see Chapter 11) described fully in Equations 14.13–14.18 and summarized in Equation 14.19. To begin with, whether a species can be present or not (wi in Table 14.5) is assumed to be an independent random variable, and not dependent on the presence or absence of other species.6 Because it can take on values of either 1 or 0, wi can be modeled as a Bernoulli random variable: wi ~ Bernoulli(Ω)

(14.13)

5 This upper bound on species richness must be identified independently of the sample data, from sources such as regional checklists of species or collated museum records (Ellison et al. 2012). 6

Null models and randomization tests of unreplicated incidence matrices have traditionally been used for the analysis of species co-occurrence (Gotelli and Graves 1996). However, with replicated hierarchical sampling, these problems are beginning to be studied with occupancy models (see Chapter 8 in MacKenzie et al. 2006).

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TABLE 14.5 The complete matrix for estimating species richness Species i

Abundance

Incidence

y11

y12

…

y1s

z11

z12

…

z1s

y21

y22

…

y2s

z21

z22

…

z2s

....

yn1

yn2

…

yns

zn1

zn2

…

zns

n+1

…

zn+1,1

z n+1,2

…

z n+1,s

wn+1

n+2

…

z n+2,1

z n+2,2

…

z n+2,s

wn+2

....

…

zN1

zN2

…

zNs

N+1

…

zN+1,1

zN+1,2

…

zN+1,s

wN+1

N+2

…

zN+2,1

zN+2,2

…

zN+2,s

wN+2

....

…

This “augmented” matrix builds on the observed abundances of species (number of pitfall traps in which ants were collected) at each site (medium blue portion of the matrix, using data from Table 14.3) and the additional rows of zeros for abundances of species uncollected but present at each site (dark blue, from Table 14.4). Additional rows and columns augment this matrix (lightest blue): species potentially available for collecting from the regional species pool (rows N + 1 to M); an incidence matrix of zik; and a column vector of parameters w indicating whether or not a species in the regional species pool could have been collected. See Dorazio et al. (2011) for additional details.

where Ω is the probability that a species in the full N × s matrix is present and could be captured. The parameter of interest N, the total number of species in the community, is derived as: N = ∑ iM=1 w i

(14.14)

If we can estimate Ω and wi, we can solve for N. Getting there takes several steps. First, examine the N × s incidence matrix of ziks. Each zik is either a 1 (species present) or a 0 (species absent), but note that any species i that is not a possible member of the community (wi = 0) must have

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Occupancy

zik = 0 as well. In other words, zik is conditional on wi (see Chapter 1 for a refresher on conditional probabilities): zik | wi ~ Bernoulli(wiψik)

(14.15)

As above, y is occupancy, here of species i at site k. Occupancy ψik is multiplied by community membership wi because zik will be 1 with occupancy probability ψik if species i can be present in the community; if not, then zik will always be 0. Thus, if wi = 1, then P(zik = 1 | wi = 1) = ψik (the probability of occupancy). Otherwise, if wi = 0, then P(zik = 0 | wi = 0) = 1. Second, to estimate y, which is a function of some set of site- or timespecific covariates, we use Equation 14.11a. Recall from Equation 14.5, however, that the estimates of ψik are also dependent on detection probability pik. If we assume that each of the ith species of ant present at the kth site has equal probability of being captured in any of the Jk pitfall traps at the two sampling dates (J = 1, …, 50), then their probability of captures (= detection) in a pitfall trap can be modeled as: yik | zik ~ Binomial(Jk, zikpik)

(14.16)

Once again, note that pik is a conditional probability of capture, given that the species is there. If it is absent from the site,

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