Centrarchid Fishes
Centrarchid Fishes Diversity, Biology, and Conservation
Edited by S. J. Cooke and D. P. Philipp
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Centrarchid Fishes
Centrarchid Fishes Diversity, Biology, and Conservation
Edited by S. J. Cooke and D. P. Philipp
A John Wiley & Sons, Ltd., Publication
This edition first published 2009 2009 Blackwell Publishing Ltd. Wiley-Blackwell is an imprint of John Wiley & Sons, formed by the merger of Wiley’s global Scientific, Technical and Medical business with Blackwell Publishing. Registered office John Wiley & Sons Ltd, The Atrium, Southern Gate, Chichester, West Sussex, PO19 8SQ, United Kingdom Editorial office John Wiley & Sons Ltd, The Atrium, Southern Gate, Chichester, West Sussex, PO19 8SQ, United Kingdom For details of our global editorial offices, for customer services and for information about how to apply for permission to reuse the copyright material in this book please see our website at www.wiley.com/wiley-blackwell. The right of the author to be identified as the author of this work has been asserted in accordance with the Copyright, Designs and Patents Act 1988. All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, electronic, mechanical, photocopying, recording or otherwise, except as permitted by the UK Copyright, Designs and Patents Act 1988, without the prior permission of the publisher. Wiley also publishes its books in a variety of electronic formats. Some content that appears in print may not be available in electronic books. Designations used by companies to distinguish their products are often claimed as trademarks. All brand names and product names used in this book are trade names, service marks, trademarks or registered trademarks of their respective owners. The publisher is not associated with any product or vendor mentioned in this book. This publication is designed to provide accurate and authoritative information in regard to the subject matter covered. It is sold on the understanding that the publisher is not engaged in rendering professional services. If professional advice or other expert assistance is required, the services of a competent professional should be sought. Library of Congress Cataloging-in-Publication Data is available A catalogue record for this book is available from the British Library. Set in 9/11 Times-Roman by Laserwords Private Limited, Chennai, India Printed and bound in Singapore by Fabulous Printers Pte Ltd 1 2009
Contents
List of Contributors Acknowledgments Preface About the Editors Chapter 1
Chapter 2
Chapter 3
xi xiii xv xvii
Species diversity, phylogeny and phylogeography of Centrarchidae T. J. Near and J. B. Koppelman
1
1.1 Introduction
1
1.2 Species diversity
1
1.3 Centrarchid fossils
5
1.4 Phylogeny
12
1.5 Phylogeography
26
1.6 Conclusions and future directions
30
1.7 Acknowledgments
31
References
31
Hybridization and speciation in centrarchids D. I. Bolnick
39
2.1 Introduction
39
2.2 Incidence of hybridization in centrarchids
39
2.3 What centrarchid hybrids tell us about speciation
41
2.4 Applied value of hybrids
58
2.5 Hybrids as a conservation threat
60
2.6 Future directions
61
2.7 Conclusions and summary
62
References
62
Ecomorphology of centrarchid fishes D. C. Collar and P. C. Wainwright
70
3.1 Introduction
70
3.2 Ecomorphology of feeding
71
3.3 Ecomorphology of locomotion
81
3.4 Conclusions
85
References
85 v
vi
Contents
Chapter 4
Chapter 5
Chapter 6
Chapter 7
Alternative reproductive tactics in the Centrarchidae B. D. Neff and R. Knapp
90
4.1 Introduction
90
4.2 Alternative reproductive tactics in the Centrarchidae
92
4.3 Genetic mechanisms for alternative reproductive tactics
94
4.4 Proximate mechanisms for alternative reproductive tactics
97
4.5 Ecological and evolutionary constraints on the evolution of alternative reproductive tactics
98
4.6 Alternative reproductive tactics in other fishes
99
4.7 Future directions
100
References
100
Early life history and recruitment D. R. DeVries, J. E. Garvey, and R. A. Wright
105
5.1 Introduction
105
5.2 Definition of “early life history” and “recruitment”
105
5.3 Variation in early life history across the centrarchids
105
5.4 Meta-analysis of life history data for several centrarchids
108
5.5 Recruitment in the centrarchids
113
5.6 Some general findings from the literature review
121
5.7 Search for critical periods
121
5.8 Evidence for broad groupings within the Centrarchidae
122
5.9 Gaps in our knowledge/research and management needs
123
References
123
Population and community ecology of Centrarchidae D. D. Aday, J. J. Parkos III, and D. H. Wahl
134
6.1 Introduction
134
6.2 Population ecology of Lepomis
134
6.3 Micropterus
140
6.4 Other centrarchids
144
6.5 Community ecology
148
6.6 Conclusions
154
6.7 Current and future directions
154
References
155
Centrarchid energetics M. S. Bevelhimer and J. E. Breck
165
7.1 Introduction
165
7.2 Centrarchid bioenergetics models
166
7.3 Food consumption and feeding energetics
171
Contents
7.4 Metabolic rate
Chapter 8
Chapter 9
vii
176
7.5 Energetic wastes (egestion, excretion, and SDA)
184
7.6 Growth energetics
184
7.7 Reproductive energetics
191
7.8 Synthesis
195
7.9 Research needs
196
References
197
Physiology and organismal performance of centrarchids J. D. Kieffer and S. J. Cooke
207
8.1 Introduction
207
8.2 Baseline physiological variables
208
8.3 Physiological challenges/tolerances
208
8.4 Physiological response to stress in centrarchids
222
8.5 Cardiovascular physiology
228
8.6 Thermal biology
242
8.7 Conclusions
250
References
251
Winter biology of centrarchid fishes C. D. Suski and M. S. Ridgway
264
9.1 Introduction
264
9.2 Definition of “winter”
264
9.3 Current research
264
9.4 Temperature
265
9.5 Dissolved oxygen and winterkill
267
9.6 Physical and physiological changes
268
9.7 Swimming abilities
269
9.8 Species ranges and life history traits
270
9.9 General activity level
271
9.10 Winter movements
273
9.11 Feeding
274
9.12 Growth
276
9.13 Aggregations
277
9.14 Winter habitat
278
9.15 Photoperiod
279
9.16 Overwinter survival
279
9.17 Conclusions and future directions
282
References
284
viii
Contents
Chapter 10
Chapter 11
Chapter 12
Centrarchid aquaculture J. E. Morris and R. D. Clayton
293
10.1 Introduction
293
10.2 Historical review
293
10.3 Culture facilities
294
10.4 Lepomis culture (bluegills and their hybrids)
295
10.5 Pomoxis spp. culture
300
10.6 Micropterus spp. culture
302
10.7 Future for centrarchids as aquaculture species
305
References
307
Centrarchid fisheries S. Quinn and C. Paukert
312
11.1 Introduction
312
11.2 Historical fisheries
312
11.3 Recreational fisheries for black bass Micropterus spp.
317
11.4 Recreational fisheries for crappie Pomoxis spp.
319
11.5 Recreational fisheries for sunfish Lepomis spp.
320
11.6 Fisheries for Ambloplites spp.
322
11.7 Recreational fisheries for other centrarchids
323
11.8 Regulations
323
11.9 Future considerations in centrarchid management
326
References
330
Contemporary issues in centrarchid conservation and management S. J. Cooke, K. C. Hanson, and C. D. Suski
340
12.1 Introduction
340
12.2 Threats to centrarchid fishes and strategies for minimizing threats
340
12.3 Introduction of exotics
347
12.4 Environmental alteration and degradation
349
12.5 Stocking—mixing of populations and outbreeding
355
12.6 Parasites and diseases
355
12.7 Exotic centrarchids as threats to conservation
356
12.8 Global conservation status of centrarchids
358
12.9 Conclusion
359
References
359
Contents
Chapter 13
ix
Centrarchid identification and natural history M. L. Warren, Jr.
375
13.1 Introduction
375
13.2 Generic and species accounts
377
13.3 Acantharchus pomotis (Baird)
377
13.4 Ambloplites Rafinesque
379
13.5 Archoplites interruptus (Girard)
389
13.6 Centrarchus macropterus (Lac´ep`ede)
392
13.7 Enneacanthus Gill
393
13.8 Lepomis Rafinesque
400
13.9 Micropterus Lac´ep`ede
434
13.10 Pomoxis Rafinesque
468
13.11 Identification keys to genera and species
475
References
482
Index Color plate (between pages 334 and 335)
535
List of Contributors
D. D. Aday Department of Zoology, North Carolina State University, Raleigh, North Carolina, USA M. S. Bevelhimer Environmental Sciences Division, Oak Ridge National Laboratory, Oak Ridge, Tennessee, USA D. I. Bolnick Section of Integrative Biology, University of Texas at Austin, Austin, Texas, USA J. E. Breck Institute for Fisheries Research, Michigan Department of Natural Resources and University of Michigan, Ann Arbor, Michigan, USA R. D. Clayton Department of Natural Resources and Environmental Management, Iowa State University, Ames, Iowa, USA D. C. Collar Section of Evolution and Ecology, University of California, Davis, California, USA S. J. Cooke Fish Ecology and Conservation Physiology Laboratory, Department of Biology and Institute of Environmental Science, Carleton University, Ottawa, Ontario, Canada D. R. DeVries Department of Fisheries and Allied Aquacultures, Auburn University, Auburn, Alabama, USA J. E. Garvey Fisheries and Illinois Aquaculture Center and Department of Zoology, Southern Illinois University, Carbondale, Illinois, USA K. C. Hanson Fish Ecology and Conservation Physiology Laboratory, Department of Biology, Carleton University, Ottawa, Ontario, Canada J. D. Kieffer Department of Biology and Canadian Rivers Institute, University of New Brunswick, Saint John, New Brunswick, Canada R. Knapp Department of Zoology, University of Oklahoma, Norman, Oklahoma, USA J. B. Koppelman Missouri Department of Conservation, Columbia, Missouri, USA J. E. Morris Department of Natural Resources and Environmental Management, Iowa State University, Ames, Iowa, USA T. J. Near Department of Ecology and Evolutionary Biology and Peabody Museum of Natural History, Yale University, New Haven, Connecticut, USA B. D. Neff Department of Biology, University of Western Ontario, London, Ontario, Canada J. J. Parkos III Illinois Natural History Survey, Division of Ecology and Conservation Sciences, Champaign, Illinois, USA C. Paukert United States Geological Survey, Kansas Cooperative Fish and Wildlife Research Unit, Division of Biology, Kansas State University, Manhattan, Kansas, USA xi
xii
List of Contributors
D. P. Philipp Illinois Natural History Survey, Division of Ecology and Conservation Sciences, Champaign, Illinois, USA S. Quinn In-Fisherman Incorporated, Baxter, Minnesota, USA M. S. Ridgway Harkness Laboratory of Fisheries Research, Aquatic Research and Development Section, Ontario Ministry of Natural Resources and Trent University, Peterborough, Ontario, Canada C. D. Suski Department of Natural Resources and Environmental Sciences, University of Illinois, Urbana, Illinois, USA D. H. Wahl Illinois Natural History Survey, Kaskaskia Biological Station, Sullivan, Illinois, USA P. C. Wainwright Section of Evolution and Ecology, University of California, Davis, California, USA M. L. Warren, Jr. Center for Bottomland Hardwoods Research, Southern Research Station, United States Department of Agriculture Forest Service, Mississippi, USA R. A. Wright Department of Fisheries and Allied Aquacultures, Auburn University, Auburn, Alabama, USA
Acknowledgments
T. J. Near and J. B. Koppelman (Chapter 1) thank several anonymous reviewers. J. B. Koppelman was supported by the Missouri Department of Conservation. D. I. Bolnick (Chapter 2) thanks S. J. Cooke, T. J. Near, D. P. Philipp, and two anonymous reviewers for comments on early drafts of this chapter. Research for this chapter was supported by NSF grant #DEB-0412802 to D. I. Bolnick, and the University of Texas at Austin. D. C. Collar and P. C. Wainwright (Chapter 3) thank two anonymous reviewers whose comments improved their chapter. They are also grateful to D. I. Bolnick, A. Carroll, S. Day, T. Higham, and T. J. Near for their insights into centrarchid biology. D. C. Collar was supported by a U.C. Davis Center for Population Biology fellowship and NSF Grant # IOB-0444554 to P. C. Wainwright. B. D. Neff and R. Knapp (Chapter 4) thank M. R. Gross and N. Santangelo for comments on the chapter. B. D. Neff was supported by the Natural Sciences and Engineering Research Council of Canada and R. Knapp was supported by the National Science Foundation (IBN 0349449) and a University of Oklahoma Presidential International Travel Fellowship. D. R. DeVries, J. E. Garvey, and R. A. Wright (Chapter 5) would like to acknowledge the Alabama Department of Conservation and Natural Resources, Auburn University’s Department of Fisheries and Allied Aquacultures, and the Southern Illinois University at Carbondale Fisheries and Illinois Aquaculture Center for their support while they worked on this book chapter. D. D. Aday, J. J. Parkos III, and D. H. Wahl (Chapter 6) wish to thank M. Carey and L. Einfalt, both of the Illinois Natural History Survey, who assisted with the literature review and preparation of figures. This chapter was improved by the thoughtful reviews of J. R. Jackson, B. R. Robinson, and S. J. Cooke. M. S. Bevelhimer and J. E. Breck (Chapter 7) thank several anonymous reviewers. M. S. Bevelhimer works with the Oak Ridge National Laboratory, which is managed by UT-Battelle, LLC, for the US Department of Energy under contract DE-AC05–00OR22725. The work by J. E. Breck on this chapter was funded in part by Federal Aid in Sport Fish Restoration (Dingell-Johnson), Project F-80-R, and the Fish and Game Fund of the State of Michigan. J. D. Kieffer and S. J. Cooke (Chapter 8) thank J. Schreer, C. Kieffer, S. Peake, and A. Kolok for thoughtful reviews and L. Arsenault for help with preparing figures. J. D. Kieffer thanks the University of New Brunswick (Saint John) and the MADSAM fish group for their continued support. Original research reported here by J. D. Kieffer was supported by a grant from the Natural Sciences and Engineering Research Council of Canada. S. J. Cooke was supported by the Natural Sciences and Engineering Research Council of Canada, the Illinois Natural History Survey, the University of British Columbia, and Carleton University. C. D. Suski and M. S. Ridgway (Chapter 9) thank A. Danylchuk and several anonymous reviewers for comments on an earlier draft of the manuscript. C. D. Suski was supported by the Natural Sciences and Engineering Research Council of Canada, Queen’s University, the Ontario Ministry of Natural Resources, and the University of Illinois. M. S. Ridgway was supported by the Ontario Ministry of Natural Resources. J. E. Morris and R. D. Clayton (Chapter 10) thank the Department of Natural Resources and Environmental Management at Iowa State University and the North Central Regional Aquaculture Center. S. Quinn and C. Paukert (Chapter 11) thank D. Willis for thoughtful discussions on centrarchid fisheries, and acknowledge helpful editorial suggestions from S. J. Cooke, W. Wegman, and an anonymous reviewer. This work was supported by In-Fisherman, Inc. S. J. Cooke, K. C. Hanson, and C. D. Suski (Chapter 12) thank L. Thompson for help with final preparation of their chapter. S. J. Cooke was supported by the Natural Sciences and Engineering Research Council of Canada, an Izaak Walton Killam Fellowship from the University of British Columbia, and Carleton University. C. D. Suski was supported by the Ontario Ministry of Natural Resources, the Natural Sciences and Engineering Research Council of Canada, and the University of Illinois. xiii
xiv
Acknowledgments
M. L. Warren, Jr. (Chapter 13) thanks T. Darden (Enneacanthus), C. S. Schieble (Ambloplites), and P. Crain, R. Schwartz, and C. M. Woodley (Archoplites) for constructive reviews of drafts of the chapter. For sharing ongoing research, alerting him to information sources, or numerous other courtesies, he gratefully acknowledges B. M. Burr, R. C. Cashner, A. C. Commens-Carson, K. S. Cummings, T. Darden, B. Fisher, W. R. Haag, R. E. Jenkins, A. E. Keller, P. Crain, R. M. Mayden, J. G. McWhirter, M. O’Connell, K. Oswald, L. M. Page, M. S. Peterson, F. C. Rohde, C. S. Schieble, R. Schwartz, W. C. Starnes, and C. M. Woodley. L. Thompson kindly formatted the references for this chapter.
Preface
The fishes in the family Centrarchidae are more commonly known as the freshwater sunfishes, a warmwater clade with 34 extant species that are endemic to North America. This group of warmwater fishes generally consists of small to moderately sized individuals that are highly colored (like the bluegill on the cover of the book). The sunfish family includes such prominent sportfish species as the largemouth bass, Florida bass, smallmouth bass, and bluegill. The largemouth bass is the most popular recreational sportfish in North America and is the basis for a large industry. In addition, bass are the frequent quarry of anglers participating in competitive angling events. Centrarchid fishes also play important ecological roles in structuring communities. They are commonly the dominant top-level predators in the diverse lentic and lotic warmwater communities of freshwater fishes in eastern North America. They provide forage for many other species and also serve as hosts for sensitive life-stages of threatened bivalves. The reproductive strategies of centrarchid fishes are especially interesting in that the male fish provide sole parental care for offspring over periods ranging from maybe as short as 1 or 2 days (for Sacramento perch) to 4 to 5 weeks (for smallmouth bass). In addition, centrarchids have been widely introduced around the globe, leading to a number of conservation concerns. Due to the popularity of this group of fishes, state and provincial fisheries managers devote substantial efforts toward managing these species. Some regions place significant emphasis on stock enhancement using cultured fish. Although there has been a recent explosion of research on sunfish species in response to their abundance and importance, at present, this large body of literature is not supported by any comprehensive syntheses on the biology and ecology of these fishes. As managers and scientists press forward with research, management, and conservation strategies, there will be an increased need to coalesce the disparate accounts of sunfish biology. Indeed, an understanding of their physiology and behavior is essential for understanding the magnitude of the threats faced by these fishes. The tome that we have developed with our team of expert authors represents a synthesis of the current state of knowledge on sunfish biology. An overriding goal of the book is to celebrate the life-history variation evident in this group of fishes (hence the use of the word “diversity” in the title of the book). A secondary objective was to summarize the linkages between basic ecology and the applied management and conservation of centrarchid fishes. Contributors were asked both to synthesize the existing literature and to contribute novel data from unpublished or forthcoming works. For that reason, we developed a team of contributors that represent those individuals at the cutting edge of centrarchid research. Authors were asked to provide coverage of all species, not just those of economic importance. Almost every author, however, identified that the majority of the available data and research were focused on several species (largemouth bass, smallmouth bass, bluegill, pumpkinseed, rock bass, black crappie, and white crappie). Detailed species accounts (of 33 of the 34 extant species; excludes the Alabama bass, Micropterus henshalli ) and a key to the centrarchids were developed by M. Warren and can be found at the end of the book. In total, the book contains 13 chapters that cover almost all aspects of sunfish biology and management. One notable omission from the list of chapters is the one focused on the reproductive biology of the centrarchid fishes. This is intentional because various aspects of reproduction were included in all of the chapters (e.g., hybridization, early life history, population biology, energetics, culture) and are thus covered throughout the book. Alternative reproductive tactics are covered independently. The detailed species accounts at the end of the book also include summaries of reproductive biology for all of the centrarchid fishes. We are particularly excited to include a chapter on winter biology, a topic of high importance to centrarchid fishes (i.e. overwinter mortality can influence recruitment), particularly toward the northern edge of their range. To our knowledge, there are no other “taxon” specific tomes that include coverage of winter biology. Thanks to the talented authors and the many referees, we are confident that this book is THE stand-alone reference on the biology of one of the most important groups of fishes in North America. Emphasizing the diversity of centrarchid fishes and the ongoing research efforts to clarify phylogenetic relationships, during the period when the book was being typeset an additional centrarchid was elevated to the species level. That addition is Micropterus henshalli (Hubbs and Bailey 1940), the Alabama Bass (See Baker, W. H., C. E. Johnston, and xv
xvi
Preface
G. W. Folkerts, 2008. The Alabama Bass, Micropterus henshalli (Teleostei: Centrarchidae), from the Mobile River basin. Zootaxa 1861:57–67). The previously recognized subspecies of Micropterus punctulatus from the Mobile River system of Alabama, Georgia, and Mississippi has been elevated to species status on the basis of morphological evidence, but it had long been recognized as distinct according to ecological, morphometric/meristic, and genetic characters. As such, although we formally recognize 34 extant centrarchid species, only 33 of them are covered extensively in this book. We do not include a formal natural history account for Micropterus henshalli, however, details can be found under the account for Micropterus punctulatus where it is described as a subspecies. We were able to make some limited changes at the proof stage to Chapter 1 in recognition of this taxonomic change, however, the phylogenies presented exclude this species. Given the many advances in molecular genetics and taxonomy, we would expect that the number of centrarchid fish species would increase in the coming years. Hence, although an inconvenience to those of us working on this book project, this taxonomic elevation is further evidence of the diversity of centrarchid fishes. We thank the many individuals that contributed to the book either intellectually or in the form of other support. This project was initiated when Cooke was an NSERC and Killam Post-Doctoral Fellow in the Centre for Applied Conservation Research at the University of British Columbia. At the time, Cooke was mentored by Dr. S. Hinch and Dr. T. Farrell, both of whom provided the freedom and encouragement to pursue this project. In the final phases of editing, Cooke was supported by the Natural Sciences and Engineering Research Council of Canada, the Rainy Lake Fisheries Charity Trust, the Ontario Ministry of Research and Innovation (Early Researcher Award), and Carleton University. D. Philipp was supported by the Illinois Natural History Survey and the Illinois Department of Natural Resources. We recognize and appreciate tremendously that the Queen’s University Biological Station provided a stimulating, productive, and fun environment to launch the idea for the book. We are particularly indebted to A. Weckworth and L. Thompson who completed detailed technical editing to ensure consistency in format and style throughout the book. D. Ramesh, Project Manager from Laserwords Private Limited in India, provided additional technical editing and facilitated the typesetting and proof changes during the final phases of the publication process. We also thank our families for continued support and acceptance of our crazy field schedules. From Blackwell Science Publishers (UK), N. Balmforth, L. Price, and K. Nuttall provided support and continual encouragement throughout the protracted writing and editing process. We also wish to acknowledge all of the authors for providing contributions that were of high quality and incredibly comprehensive. The project took several years to complete, and our authors were extremely patient. Furthermore, we thank the many anonymous (unless declared and listed in the acknowledgments) referees for providing thoughtful reviews of the lengthy chapters. S. J. Cooke and D. P. Philipp, Eleuthera, The Bahamas, December 2, 2007
About the Editors
S. J. Cooke: Cooke received his undergraduate and M.Sc. degrees from the University of Waterloo. He completed his Ph.D. research at the University of Illinois in 2002 while working with Dr. D. P. Philipp and Dr. D. H. Wahl at the Illinois Natural History Survey. Cooke was then awarded an NSERC Post Doctoral Fellowship and Izaak Walton Killam Fellowship, which he held as a postdoctoral fellow at the University of British Columbia where he worked with Dr. S. Hinch and Dr. T. Farrell. In 2005, Cooke became an Assistant Professor in Environmental Science and Biology at Carleton University (Ottawa, Canada) where he is Director of the Fish Ecology and Conservation Physiology Laboratory. Cooke, his students, and collaborators, study how fish respond to natural and anthropogenic stressors and how individuals, populations, and species vary in their response. Cooke has published over 100 peer reviewed papers, about half on fish in the sunfish family covering topics such as the energetics of parental care, the physiological consequences of angling practices, and the consequences of outbreeding on fish performance. Cooke has been the recipient of the American Fisheries Society Award of Excellence in Fisheries Management and an Early Researcher Award from the Ontario Ministry of Innovation. He is also an editor for the journal Endangered Species Research and is on the Editorial Board for Fisheries Research. Cooke is an Affiliate Scientist with the Illinois Natural History Survey, Adjunct Professor at Queen’s University, and an Honorary Research Associate at the University of British Columbia. He and his wife currently reside in Ottawa, a region rich with centrarchid dominated fisheries. D. P. Philipp: Philipp received his undergraduate degree from Lafayette College and his Ph.D. from the University of Massachusetts in 1976. He is currently Principal Scientist at the Illinois Natural History Survey (INHS) and is a Professor in three departments at the University of Illinois at Urbana-Champaign. His research interests focus on conservation genetics and behavioral ecology with a focus on centrarchid fishes. His findings have helped to elucidate the consequences of outbreeding depression, hybridization, and fisheries exploitation on centrarchid populations. In his role at the INHS, Dr. D. P. Philipp conducts research in support of the Illinois Department of Natural Resources. He is the director of the state creel survey and responsible for assessing recreational fishery dynamics throughout Illinois. Dr. D. P. Philipp has served on a number of committees including the Independent Scientific Advisory Board of the Northwest Power and Conservation Council. He is one of the initial founders of the Fisheries Conservation Foundation (an education and outreach partner with the American Fisheries Society) and currently serves on the Foundation’s Board of Directors. He has edited several prominent books including “Black Bass Ecology and Conservation” in 2002 and has over 100 papers in peer reviewed outlets. D. P. Philipp was selected as the first touring lecturer for the Zoological Education Trust of the Canadian Society of Zoologists. He and his family reside in Champaign, Illinois, but spend the spring in Canada studying centrarchid reproduction at the Queen’s University Biology Station.
xvii
Chapter 1
Species diversity, phylogeny and phylogeography of Centrarchidae T. J. Near and J. B. Koppelman
1.1 Introduction Centrarchidae is a clade of freshwater fishes endemic to North America, a part of the world that harbors more species of freshwater fishes than any other nontropical region on Earth (Briggs 1986; Lundberg et al . 2000). Centrarchid fishes have been of interest to biologists for a long period of time because they are commonly the dominant top-level predators in the diverse communities of freshwater fishes in eastern North America, and as such, they are among the world’s most popular freshwater sport fishes (Henshall 1881; Etnier and Starnes 1993; Philipp and Ridgway 2002). Interestingly, it is only in the last 10 years or so that comparative morphological and molecular data have been used in conjunction with objective character-based methods to investigate the phylogenetic relationships of Centrarchidae. The goal of this chapter is to review and assess previous ideas regarding the diversity and relationships of centrarchid species. We hope to provide biologists from all disciplines with a clear picture of the current and best-supported hypotheses of centrarchid phylogeny, and we intend to illustrate how many recent, cutting-edge efforts have agreed remarkably with studies published as far back as the nineteenth century. Although we realize our esoteric interests in centrarchid diversity and phylogeny, as well as our desire to understand the results of modern phylogenetic analyses in the context of the rich past of centrarchid taxonomy and systematics may be confusing to the average fish biologist or ichthyologist, we will attempt to clarify what seems like a morass of trees and classifications for biologists in need of phylogenetic hypotheses. It is our desire that both comparative biologists and conservation agencies exploit the current state of knowledge regarding centrarchid diversity and phylogenetic relationships. In this chapter we provide a discussion of the currently recognized diversity of both extant and fossil species in Centrarchidae, and we attempt to illuminate some unresolved issues in this area that need attention in future research efforts. We present an overview of previous investigations and hypotheses concerning the evolutionary relationships of Centrarchidae, including a discussion of recent efforts using morphological and molecular data in addition to those that pre-date the development of phylogenetic systematics, or cladistics (Hennig 1966). Many of the pre-cladistic ideas of centrarchid relationships discussed in this review were presented as purely taxonomic hypotheses, where the hypothesized relationships were implied from the composition and ranking of taxa. Evolutionary biologists often investigate genetic variation within a geographic context, as intraspecific gene trees often show a strong geographic pattern. Such is the science of phylogeography (Avise 2000). We provide a review and discussion of phylogeography in centrarchids, highlighting some of the problems that have made such analyses in Centrarchidae less straightforward than in species from other groups of North American freshwater fishes.
1.2 Species diversity 1.2.1 Extant species and the status of subspecies Currently, 34 extant species are recognized in Centrarchidae (Table 1.1), with the most recently described species being Ambloplites constellatus and Micropterus cataractae (Cashner and Suttkus 1977; Williams and Burgess 1999). As in 1
2
Centrarchid fishes
Table 1.1 Currently recognized centrarchid species and proposed classification. Fossil genera and species are indicated with a dagger. Centrarchidae (44 species: 33 extant, 11 extinct) Centrarchinae Acantharchus pomotis (Baird 1855) Mud sunfish Ambloplites ariommus (Viosca 1936) Shadow bass Ambloplites cavifrons (Cope 1868) Roanoke bass Ambloplites constellatus (Cashner and Suttkus 1977) Ozark bass Ambloplites ruprestris (Rafinesque 1817) Rockbass Archoplites †clarki (Smith and Miller 1985) Clarkia perch Archoplites interruptus (Girard 1854) Sacramento perch Archoplites †molarus (Smith et al. 2000) Ringold sunfish Archoplites †taylori (Miller and Smith 1967) Lake Idaho sunfish †Boreocentrarchus smithi (Schlaikjer 1937) Healy Creek sunfish ` 1801) Flier Centrarchus macropterus (Lacepede Enneacanthus chaetodon (Baird 1855) Blackbanded sunfish Enneacanthus gloriosus (Holbrook 1855) Bluespotted sunfish Enneacanthus obesus (Girard 1854) Banded sunfish †Plioplarchus septemspinosus (Cope 1889) John Day sunfish †Plioplarchus sexspinosus (Cope 1883) Sentinel Butte sunfish †Plioplarchus whitei (Cope 1883) Laramie sunfish Pomoxis annularis (Rafinesque 1818) White crappie Pomoxis †lanei (Hibbard 1936) Ogallala crappie Pomoxis nigromaculatus (Lesueur 1829) Black crappie Lepominae Lepomis auritus (L 1758) Redbreast sunfish Lepomis cyanellus (Rafinesque 1819) Green sunfish Lepomis gibbosus (L 1758) Pumpkinseed Lepomis gulosus (Cuvier 1829) Warmouth Lepomis humilis (Girard 1858) Orangespotted sunfish Lepomis †kansasensis (Hibbard 1936) Rhino Hill sunfish Lepomis macrochirus (Rafinesque 1819) Bluegill Lepomis marginatus (Holbrook 1855) Dollar sunfish Lepomis megalotis (Rafinesque 1820) Longear sunfish ¨ Lepomis microlophus (Gunther 1859) Redear sunfish Lepomis miniatus (Jordan 1877) Redspotted sunfish Lepomis peltastes (Cope 1870) Northern longear sunfish Lepomis punctatus (Valenciennes 1831) Spotted sunfish Lepomis †serratus (Smith and Lundberg 1972) Keigh sunfish Lepomis symmetricus (Forbes 1883) Bantam sunfish (continued)
Species diversity, phylogeny and phylogeography of Centrarchidae
3
Table 1.1 (continued). Centrarchidae (44 species: 33 extant, 11 extinct) Micropterinae Micropterus cataractae (Williams and Burgess 1999) shoal bass Micropterus coosae (Hubbs and Bailey 1940) Redeye bass ` Micropterus dolomieu (Lacepede 1802) smallmouth bass Micropterus floridanus (LeSueur 1822) Florida bass Micropterus henshalli (Hubbs and Bailey 1940) Micropterus notius (Bailey and Hubbs 1949), Suwannee bass Micropterus punctulatus (Rafinesque 1819) spotted bass Micropterus †relictus (Cavender and Smith 1975) Chapala bass ` 1802) largemouth bass Micropterus salmoides (Lacepede Micropterus treculi (Vaillant and Bocourt 1874) Guadalupe bass
many groups of animals, there are many more scientific names available than there are recognized species. Not including the names of valid extant species (Table 1.1), there are 118 nominal names that are considered synonyms for species in Centrarchidae. Of these, 11 were either new names for subspecies or were introduced as species names and have been used at some point to designate subspecies (Gilbert 1998). Of the 118 nominal names, 9 are based on hybrid centrarchids; all but 1 of these are the hybrid combinations of Lepomis cyanellus * L. macrochirus and L. cyanellus * L. gibbosus (Hubbs 1920; Hubbs and Hubbs 1932; Gilbert 1998). The contemporary view of species diversity in Centrarchidae was fairly well settled by the turn of the nineteenth and twentieth centuries, as the vast majority of valid centrarchid species were described between 1800 and 1883 (Table 1.1; Figure 1.1). This period was also when most of the synonymous names were introduced (Bailey 1938; Gilbert 1998). Through both the nineteenth and twentieth centuries centrarchid species have been described using very similar types of data from external morphology, including meristics (scale row and fin element counts), morphometrics (body proportions), pigmentation patterns, and coloration (Cope 1868, 1870; Hubbs and Bailey 1940; Cashner and Suttkus 1977; Williams and Burgess 1999). To date, comparative phylogenetic methods, using either morphological or molecular data, have not been used in describing new centrarchid species.
No. of valid species described
35 30 25 20 15 Ambloplites rupestris
10 5
1750
1800
1850
1900
1950
2000
Year described
Figure 1.1 Plot illustrating the growth of valid extant centrarchid species descriptions from the nineteenth through twentieth centuries. Ambloplites rupestris redrawn from Forbes and Richardson (1920).
4
Centrarchid fishes
The status of subspecies in Centrarchidae is much less resolved when compared to the 33 recognized valid extant species (Table 1.1). The use of subspecies in North American fish taxonomy has a relatively inconsistent history, and since the initial critique of subspecies, most modern workers in systematics have been moving away from using this rank (Wilson and Brown 1953; Burbrink et al . 2000). However, there remain 11 names that have been historically designated as centrarchid subspecies. We are able to categorize each of these names into three classes: (i) subspecies that do not exhibit significant variation from the nominal subspecies, (ii) subspecies that are based on hybrid specimens, and (iii) subspecies that merit elevation to species. Three centrarchid subspecies have been invalidated as it was demonstrated that they did not differ appreciably from other populations of the nominal species. Acantharchus pomotis mizelli Fowler and Enneacanthus chaetodon elizabethae were both described as subspecies in the 1940s based on six or seven specimens (Bailey 1941; Fowler 1945). In both cases, subsequent analyses that included many more specimens failed to reveal geographic variation consistent with the recognition of the subspecies proposed for each of these species (Sweeney 1972; Cashner et al . 1989). A similar situation exists for the Neosho Smallmouth Bass, Micropterus dolomieu velox Hubbs and Bailey. This subspecies was described based primarily on slight differences in counts of the second dorsal fin rays, pigmentation patterns, and dentition on the tongue (Hubbs and Bailey 1940). The validity of M. d. velox was subsequently dismissed on the basis of slight morphological differences and clinal gradation into the nominal M. dolomieu (Bailey 1956; Gilbert 1998), a conclusion supported by more recent analyses of nuclear gene encoded allozymes and mitochondrial DNA (mtDNA) sequence data (Stark and Echelle 1998; Kassler et al . 2002). At least one centrarchid subspecies has turned out to be based on hybrid specimens. Micropterus punctulatus wichitae Hubbs and Bailey was described as a subspecies from the Wichita Mountains of southwestern Oklahoma based on differences in scale row counts (Hubbs and Bailey 1940). However, this population was initially described as hybrids of M. punctulatus and M. dolomieu (Hubbs and Ortenburger 1929). Morphological data from M. p. punctulatus, M. p. wichitae, and M. dolomieu and historical records of nonnative M. dolomieu introductions near the type locality of M. p. wichitae support the hypothesis that this subspecies is based on hybrid M. punctulatus * M. dolomieu specimens (Cofer 1995). Genetic analysis of both nuclear and mtDNA in M. punctulatus populations from the Red and Arkansas River Basins did not reveal genetic divergence of the Wichita Mountain populations of M. punctulatus (Coughlin et al . 2003). Lepomis megalotis and L. macrochirus are two centrarchid species that are thought to be polytypic and contain described subspecies (Mayden et al . 1992; Gilbert 1998). Future research documenting morphological and genetic variation in these two complexes has the strong possibility to result in the recognition of additional valid centrarchid species. L. megalotis has four, and possibly seven, valid subspecies, L. m. megalotis (Rafinesque), L. m. aquilensis (Baird and Girard), L. m. breviceps (Baird and Girard), and L. m. occidentalis Meek (Bailey 1938). In addition, L. m. convexifrons (Baird and Girard), L. m. fallax (Baird and Girard), and L. m. popeii (Girard) are three additional forms from Texas that may represent other unrecognized species related to L. megalotis (Gilbert 1998). Unfortunately, there is no published analysis of morphological variation among these subspecies, but a Ph.D. dissertation had detected substantial morphometric variation among four of the described subspecies (Barlow 1980). An analysis of allozyme variation detected appreciable genetic divergence of L. m. breviceps and L. m. aquilensis relative to the other subspecies (Jennings and Philipp 1992). Based on morphometric and body size differences, L. peltastes Cope was elevated as a species from a subspecies of L. megalotis (Bailey et al . 2004). We suspect that several additional centrarchid species will be recognized as a result of analyses of geographic variation and phylogeny of the L. megalotis complex using comparative morphological and molecular data. There is a degree of uncertainty as to how many subspecies of Lepomis macrochirus are recognized. The problem centers on Pomotis speciosus described from Brownsville, Texas by Baird and Girard (1854). This species was subsequently synonymized with L. macrochirus by Hubbs (1935). At a later date, Hubbs and Lagler (1958) treated P. speciosus as a subspecies of L. macrochirus, concluding that the geographic range is throughout Texas and northeastern Mexico. Allozyme analyses did not detect genetic differentiation between L. m. macrochirus and L. m. speciosus (Kulzer and Greenbaum 1986), and subsequent treatments of centrarchid species diversity have not recognized L. m. speciosus (Gilbert 1998). The two valid subspecies of L. macrochirus present an interesting problem of nomenclature confusion, morphological and genetic divergence, an area of presumed secondary contact and introgression, and a biogeographic pattern and a timing of divergence seen in another centrarchid sister species pair. The nominal subspecies L. m. macrochirus Rafinesque is distributed across eastern North America except for the northern Atlantic Coast (Lee et al . 1980), while the other subspecies is endemic to the Florida Peninsula (Felley 1980). Initially, the subspecies found in Florida was designated as Lepomis
Species diversity, phylogeny and phylogeography of Centrarchidae
5
macrochirus purpurescens Cope under the premise that this subspecies extended from the Atlantic Coast of the Carolinas to the Florida Peninsula (Hubbs and Allen 1943; Hubbs and Lagler 1958). The type locality for Lepomis purpurescens is in the Yadkin River Drainage in North Carolina (Cope 1870). Subsequent morphological and molecular analyses demonstrate that this is far north of the range of the Florida subspecies (Avise and Smith 1974a; Felley 1980; Avise et al . 1984), and as Gilbert (1998) has pointed out, Cope described a Bluegill from Florida, Lepomis mystacalis (Cope 1877). Therefore, the appropriate name for the Florida Bluegill is L. macrochirus mystacalis. Lepomis m. macrochirus and L. m. mystacalis are morphologically and genetically distinct, but there is a presumed area of introgression through secondary contact along most of southern Georgia and South Carolina (Felley 1980; Avise et al . 1984). Another sister species pair in Centrarchidae, Micropterus salmoides and Micropterus floridanus, exhibit a very similar distribution and area of secondary contact and introgression (Bailey and Hubbs 1949; Philipp et al . 1983). Based on a fossil calibrated molecular phylogeny of Centrarchidae, the divergence time between M. salmoides and M. floridanus is approximately 2.8 million years ago (mya) (Near et al . 2003, 2005b). Lepomis m. macrochirus and L. m. mystacalis exhibit a very similar divergence time. We found mtDNA cytochrome b gene sequences in Genbank for five individuals of L. m. macrochirus and a single L. m. mystacalis (accession numbers: AY115975, AY115976, AY225667, AY828966, AY828967, and AY828968). The average genetic distance between these two subspecies was 4.5%, which translates to a divergence time of roughly 2.3 mya (Near et al . 2003). Future work should aim toward gathering sufficient morphological and molecular data to more precisely determine the geographic distribution of these two forms and assess if L. mystacalis is a valid species. Recently Micropterus henshalli (Hubbs and Bailey) was elevated as a valid species (Baker et al . 2008), but was long recognized as a subspecies of M. punctulatus (Hubbs and Bailey 1940). Micropterus henshalli is endemic to the Mobile Basin and there are slight morphological differences between populations above and below the Fall Line (Gilbert 1973; Baker et al . 2008). However, there are substantial differences in several meristic characters between M. henshalli and M. punctulatus (Gilbert 1973), and there are marked differences in body proportions and surprising life history and dietary differences between these two species (Gilbert 1973). Perhaps the most compelling evidence for the recognition of M. henshalli includes measures of genetic divergence and the results of phylogenetic analyses. Among 19 polymorphic allozyme loci surveyed for all Micropterus species, not a single allele was shared exclusively between M. henshalli and M. punctulatus, and a fixed unique allele was found in M. p. henshalli (Kassler, et al ., 2002). In a phylogenetic analysis of Micropterus species using gap coded continuous morphological characters M. henshalli and M. punctulatus did not form a clade (Harbaugh 1994), and these two species were sister lineages in frequency parsimony of allozyme alleles (Kassler et al . 2002). In addition, molecular phylogenetic analyses of mtDNA sequences from cytb and ND2 resulted in tree topologies where M. henshalli was nested within M. coosae and distantly related to M. punctulatus (Kassler et al . 2002). Given the evidence presented above, the classification of M. henshalli as a subspecies of M. punctulatus was not compelling and the recognition of this species is supported by the substantial comparative data.
1.3 Centrarchid fossils The fossil record of Centrarchidae is fairly rich and extends in geologic time from the Late Eocene to Early Oligocene of approximately 35 mya to the very early Holocene of approximately 10 years ago. Both extant centrarchid species and centrarchid fossils are found only in North America, indicating that origin and diversification of this clade did not involve other continental regions. There are 11 valid and extinct centrarchid species known only from fossil material (Table 1.1; Figures 1.2–1.17), and there are fossils of seven extant species. Despite an excitingly abundant centrarchid fossil record, at least four of the oldest fossil centrarchid species are generally unknown to science. These fossil species are undescribed and have been under study for at least three decades. Unfortunately, they have not been made available to other researchers for study, which has significantly hindered progress in understanding the evolutionary origin of Centrarchidae and its patterns of diversification. The meager information available for these four undescribed fossil species that we present here is from general synopses of the fossil record of North American freshwater fishes (Cavender 1986, 1998). The first of these four we call the High Plains Sunfish, from the northwestern part of Montana near the foothills of the Rocky Mountains. Cavender (1986, 1998) indicates that they are found in Late Eocene to Early Oligocene deposits, but more precise age estimates are unavailable. The High Plains Sunfish has three anal spines and an emarginate caudal fin. The second of these undescribed fossils is the Chadron Sunfish
6
Centrarchid fishes
Plioplarchus sexspinosus
Figure 1.2 Photos and drawings of fossil Centrarchidae species: †Plioplarchus sexspinosus Sentinel Butte Sunfish, photo redrawn from Eastman (1917).
Plioplarchus whitei
Figure 1.3 Photos and drawings of fossil Centrarchidae species: †Plioplarchus whitei Laramie Sunfish, redrawn from Cope (1884).
Plioplarchus septemspinosus
Figure 1.4 Photos and drawings of fossil Centrarchidae species: †Plioplarchus septemspinosus John Day Sunfish, photo redrawn from Eastman (1917).
Boreocentrarchus smithi
Figure 1.5 Photos and drawings of fossil Centrarchidae species: †Boreocentrarchus smithi Healy Creek Sunfish, redrawn from Schlaikjer (1937).
Species diversity, phylogeny and phylogeography of Centrarchidae
7
Pomoxis lanei
Figure 1.6 Photos and drawings of fossil Centrarchidae species: Pomoxis †lanei Ogallala Crappie, photo redrawn from Hibbard (1936).
Pomoxis sp.
Figure 1.7 Photos and drawings of fossil Centrarchidae species: Pomoxis †sp. Wakeeney Crappie, redrawn from Wilson (1968).
Archoplites clarkii
Figure 1.8 Photos and drawings of fossil Centrarchidae species: Archoplites †clarki Clarkia Perch, photo provided by Smith (1963).
Archoplites taylori
2 mm
2 mm
2 mm 2 mm
Figure 1.9 Photos and drawings of fossil Centrarchidae species: Archoplites †taylori Lake Idaho Sunfish, redrawn from Miller and Smith (1967).
8
Centrarchid fishes
Archoplites molarus
1 cm
1 cm
1 cm
1 cm
1 cm
Figure 1.10 Photos and drawings of fossil Centrarchidae species: Archoplites †molarus Ringold Sunfish, redrawn from Smith et al . (2000).
Lepomis kansasensis
Figure 1.11 Photos and drawings of fossil Centrarchidae species: Lepomis †kansasensis Rhino Hill Sunfish, photo redrawn from Hibbard (1936).
Lepomis serratus
2 mm
2 mm 2 mm
1 mm Figure 1.12 Photos and drawings of fossil Centrarchidae species: Lepomis †serratus Keigh Sunfish, redrawn from Smith and Lundberg (1972).
Species diversity, phylogeny and phylogeography of Centrarchidae
9
Lepomis sp. A
Figure 1.13 Photos and drawings of fossil Centrarchidae species: Lepomis †sp. A Valentine Sunfish, redrawn from Smith (1962).
Lepomis sp. B
Figure 1.14 Photos and drawings of fossil Centrarchidae species: Lepomis †sp. B Wakeeney Sunfish, redrawn from Wilson (1968).
Micropterus relictus
Figure 1.15 Photos and drawings of fossil Centrarchidae species: Micropterus †relictus 1975 Chapala Bass, redrawn from Smith et al . (1975).
Micropterus sp. B
Figure 1.16 Photos and drawings of fossil Centrarchidae species: Micropterus †sp. B Wakeeney Bass, redrawn from Wilson (1968).
Micropterus sp. C
Figure 1.17 Photos and drawings of fossil Centrarchidae species: Micropterus †sp. C Laverne Bass, redrawn from Smith (1962).
10
Centrarchid fishes
from Lower Oligocene limestone deposits in the South Dakota Badlands, dating this fossil to the White River group of approximately 28 to 35 mya (Tedford et al . 1987). The Chadron Sunfish has three anal spines and 27 to 28 vertebrae (Cavender 1986). The third fossil sunfish in this group of undescribed forms is from Lower Miocene deposits in South Dakota, and Cavender (1986) provides an age of approximately 25 mya. These are very similar in morphology to the Chadron Sunfish, but have 29 vertebrae (Cavender 1986). The last of the four undescribed fossils in Cavender (1986) is from Middle Miocene deposits, but no location is given. This fossil species has six or seven anal fin spines and is similar to fossils that were assigned to †Plioplarchus (Cope 1884). There are two extinct genera of Centrarchidae known from the fossil record, †Plioplarchus and †Boreocentrarchus. †Plioplarchus contains three species (Table 1.1), and is the oldest of the described centrarchid fossils (Figures 1.2–1.4). †Plioplarchus sexspinosus and †P. whitei were described from Oligocene age freshwater limestone deposits from the Sentinel Butte of North Dakota (Cope 1883) that date to approximately 30 mya (Feldman 1962) (Figures 1.2 and 1.3). †Plioplarchus sexspinosus and †P. whitei are also found in the Badlands of South Dakota in the White River Group. Specimens that are either †P. sexspinosus or †P. whitei are found at the contact between the Chadron and Brule Formations (Welzenbach 1992), and this is dated to approximately 31 mya (Tedford et al . 1987). †Plioplarchus septemspinosus was described from the John Day River in Oregon (Cope 1889) in the geological deposits that make up the John Day Fauna (Figure 1.4), and is dated between 18 and 31 mya (Tedford et al . 2004). Fossils currently assigned to †P. septemspinosus are also found in the Trout Creek Flora in Oregon and this is dated at 13 mya (Graham 1999). Morphological analyses indicate that †P. septemspinosus from the John Day and Trout Creek locations in Oregon are different from each other and both of these are quite divergent from †P. sexspinosus and †P. whitei (Schlaikjer 1937; Bailey 1938; Smith and Miller 1985). These differences were substantial enough for Bailey (1938) in his unpublished Ph.D. dissertation to describe a new genus for †P. septemspinosus. †Boreocentrarchus smithi was described from Healy Creek, Alaska in deposits that were thought to age from the Oligocene to the Early Miocene (Figure 1.5) (Schlaikjer 1937; Uyeno and Miller 1963), and a more precise estimate of this formation at 24 to 18 mya agrees with these earlier estimates (Merritt 1987). Schlaikjer (1937) argues that †B. smithi is closely related to †P. septemspinosus, but others have questioned whether †B smithi is a centrarchid (Uyeno and Miller 1963). Both †Plioplarchus and †Boreocentrarchus are classified in the Centrarchinae (Table 1.1), because these species possess more than three anal fin spines. Undescribed fossil species in this clade include one from the Horse Creek Fish Quarry in Laramie Co., Wyoming, that dates to approximately 19 mya (Cassiliano 1980), another from the Bear Valley, California (Smith and Miller 1985), and a third from the Humboldt Formation, Nevada, that dates to 9 mya (Smith and Miller 1985; Smith et al . 2002). The remaining centrarchid fossil species are classified in genera that also contain extant species (Table 1.1). Pomoxis is known from the fossil record with one described species, P . †lanei, and one undescribed fossil species. Pomoxis †lanei was found in the Rhino Hill Quarry in Logan Co., Kansas (Hibbard 1936), and age of this fossil formation is correlated with Coffee Ranch mammals that date to 6.6 mya (Wallace 1997; Passey et al . 2002). The holotype of P. †lanei is a complete and crushed skeleton (Figure 1.6). The specimen is a remarkable impression and many morphological features can be scored, counted, or measured (Hibbard 1936). The phylogenetic position of P. †lanei in Pomoxis is unresolved due to conflicting characters. The presence of seven dorsal fin spines and a long dorsal fin base supports the hypothesis that P. †lanei and P. nigromaculatus are sister species (Smith 1962). However, the hypothesis that P. nigromaculatus and P. annularis are sister species is supported by the presence of 17 to 20 anal fin rays in these species versus 12 anal fin rays in P. †lanei (Uyeno and Miller 1963). There is a second fossil species of Pomoxis that is undescribed. These fossils were found in the Wakeeney local fauna that is a part of the Ogallala Formation in Kansas (Wilson 1968). The age of this formation was placed in the lower portion of the Ash Hollow or upper Valentine Formation (Wilson 1968), and this dates to approximately 12 mya (Tedford et al . 2004). These are the oldest Pomoxis fossils and they are fragmentary, consisting of a dentary and premaxillary fragments (Figure 1.7). Archoplites contains three fossil species and only one extant species (Table 1.1). The oldest of the Archoplites fossil species is A. †clarki from the Clarkia Lake Beds in Idaho (Figure 1.8) (Smith and Miller 1985). This fossil formation has been dated at 15.5 mya (Golenberg et al . 1990; Wing 1998). Archoplites †taylori is found in seven different fossil locations in southwestern Idaho and these sites are characterized as lacustrine deposits (Figure 1.9). The oldest of the fossil sites containing A. †taylori is the Poison Creek formation and is dated at 9 mya (Smith and Cossel 2001). The youngest formation containing A. †taylori fossils is Jackass Butte, a part of the Grandview local fauna dated at 2.2 mya (Smith 1975; Lundelius et al . 1987). Archoplites †molarus was recently described from the Ringold Formation
Species diversity, phylogeny and phylogeography of Centrarchidae
11
in Washington (Figure 1.10). Fossils of A. †molarus are found at three different locations in the Ringold Formation and the ages of these deposits extend through the Pliocene. Fossils from the White Bluffs local fauna are the oldest at 4.5 mya, the Blufftop Locality and local fauna dates to 3.7 mya, and Tauton Locality dated at 2.9 mya contains the youngest A. †molarus fossils (Smith et al . 2000; Van Tassell et al . 2001). The oldest fossils assigned to the extant species Archoplites interruptus are from the Cache Formation in Lake Co., California and date to the Early Pleistocene, approximately 2.5 mya (Casteel and Rymer 1975). The youngest A. interruptus fossils are from Sacramento Co., California and date to the Pleistocene of approximately 100,000 years ago (Hansen and Begg 1970). There is the possibility of undescribed fossil species of Archoplites. Van Tassell et al . (2001) mention specimens from Grande Ronde Valley in Union Co., Oregon that date to 3.7 mya, and there are other Archoplites fossil specimens dated to the Early or Middle Pleistocene from Moses Lake in Washington (Miller 1965). There are four fossil species of Lepomis, and two of these are closely related to L. gulosus. Lepomis †kansasensis was found in the same fossil formation as P. †lanei, so it is dated at 6.6 mya (Hibbard 1936; Wallace 1997; Passey et al . 2002). The holotype is a nearly complete skeletal impression with a badly crushed head, but morphological features such as dentition can be distinguished (Figure 1.11). The presence of pterygoid teeth in L. †kansasensis led to the original classification of this species in Chaenobryttus that also contained L. gulosus (Hibbard 1936; Bailey 1938). Lepomis †serratus was described from fossils collected at the Keim Formation in the Sand Draw local fauna in Brown Co., Nebraska (Smith and Lundberg 1972). The age of this formation is dated at 3.4 mya (Repenning 1987). The Sand Draw L. †serratus fossils are dentaries, articulars, maxillae, prevomers and preopercles (Figure 1.12). Lepomis †serratus maxillae, prevomers, articulars, and preopercles, premaxillae, and dentaries are also reported from the Seneca local fauna in Hooker Co., Nebraska (Bennett 1979), and this is a younger fossil formation dated between 2.5 and 2.0 mya (Bell et al . 2004). Lepomis †serratus was classified in Chaenobryttus on the basis of morphological similarity of the preopercle with L. gulosus (Smith and Lundberg 1972). The initial classifications proposed for L. †kansasensis and L. †serratus indicate a fairly close phylogenetic affinity with L. gulosus, illustrated by the fact that at one point L. †kansasensis was synonymized with L. gulosus (Branson and Moore 1962). However, Smith and Lundberg (1972) argue that L. †kansasensis is more closely related to other Lepomis species than to L. gulosus, but concur that a definitive conclusion on this issue would result only from a more thorough analysis of the fossil material. There are at least two undescribed Lepomis fossil species. Both of these fossils are quite old and are represented by fragmentary material of the lower jaw. The first of these is referred to as L. †sp. A, and was initially identified as L. cf. microlophus (Smith 1962). The fossil comes from the Lower Valentine Formation in Brown Co., Nebraska (Figure 1.13) (Smith 1962). This fossil location was later identified as the Norden Bridge local fauna (Estes and Tihen 1964), and is dated at 13.5 mya (Tedford et al . 2004). The second undescribed Lepomis fossil species is referred to as L. †sp. B (Figure 1.14), and is found in the same fossil location as the undescribed Pomoxis fossil species discussed earlier (Wilson 1968), so this fossil also dates to approximately 12 mya. There are four fossil Micropterus species, but only one is described. Micropterus †relictus was described from and its jaw element fossils found in Late Pliocene–Early Pleistocene deposits in the Lake Chapala Basin, Mexico (Figure 1.15) (Smith et al . 1975). Micropterus †sp. A consists of fragmentary skull pieces and vertebrae from the Lower Snake Creek fauna in Sioux Co., Nebraska (Matthew 1924), and dates between 16 and 15 mya (Tedford et al . 2004). Micropterus †sp. B was originally identified as M. cf. punctulatus, and the fossil material is a lower pharyngeal jaw and a dentary fragment (Figure 1.16). The fossils were collected in the same formation as Pomoxis †sp. and Lepomis †sp. B and date to 12 mya (Wilson 1968). Micropterus †sp. C. is a dentary from the Laverne Formation in Beaver Co., Oklahoma (Figure 1.17) with an estimated age between 10.5 and 9.5 mya (Smith 1962; Tedford et al . 2004). In addition to A. interruptus discussed earlier, there are at least six extant centrarchid species present in the fossil record. The oldest formations that contain extant species are the Rexroad local fauna in Meade Co., Kansas and the Sand Draw local fauna in Brown Co., Nebraska (Smith 1962; Smith and Lundberg 1972), both dated at 3.4 mya (Bell et al . 2004). These two formations combined contain fossil specimens of Ambloplites rupestris, L. cyanellus, and L. humilis. Both L. cyanellus and L. humilis occur in numerous Pleistocene fossil deposits. Lepomis cyanellus is found in Pleistocene formations ranging in age from 2.3 mya to 10,000 years ago and these sites are spread across Kansas, Michigan, Nebraska, Oklahoma, and Texas (Smith 1954, 1958, 1963; Schultz 1965; Hibbard and Dalquest 1966; Lundberg 1967; Eshelman 1975; Neff 1975; Shoshani and Smith 1996). Pleistocene formations containing L. humilis fossils range in age from 2.5 mya to 250,000 years ago and are restricted to Kansas, Nebraska, and South Dakota (Smith 1963; Ossian 1973; Neff 1975; Bennett 1979; Cross et al . 1986). L. megalotis fossils are found at two Pleistocene fossil formations. The oldest of
12
Centrarchid fishes
these two locations is the Rita Blanca Lake Deposit in Hartley Co., Texas and this formation is dated at 2.4 mya (Anderson and Kirkland 1969; Koster 1969; Lindsay et al . 1975; Lundelius et al . 1987; Repenning 1987), whereas the Kanopolis local fauna in Ellsworth Co., Kansas has yielded much younger L. megalotis fossils dated at 300,000 years ago (Neff 1975; Repenning 1987). Micropterus salmoides fossils have been reported from Pleistocene deposits in Kansas, Michigan, and South Dakota that are dated from 300,000 to 14,000 years (Smith 1963; Wilson 1967; Ossian 1973; Neff 1975). Fossil L. gibbosus specimens are known only from a single Pleistocene locality in South Dakota (Ossian 1973).
1.4 Phylogeny 1.4.1 Pre-Cladistic concepts of centrarchid evolutionary relationships The nineteenth century was the time when most of the valid centrarchid species were described (Figure 1.1). Associated with this period of activity was the initial development of hypotheses of centrarchid relationships. In these early studies evolutionary relationships were reflected by the composition of species in particular taxonomic groups that were arranged in nested hierarchical ranks. For example, at the taxonomic rank of family, centrarchids were initially classified with Percidae (G¨unther 1859), implying a close relationship with pikeperches (Sander), perches (Perca), and darters (e.g., Etheostoma and Percina). The name Ichthelidae was applied to the centrarchids when they were first grouped apart from Percidae as a distinct family (Holbrook 1860). This family name was not used by later authors as Icthelis was regarded as a synonym (Bailey 1938). The first use of the name Centrarchidae came at a time when most of the valid species were described (Cope 1868). Subsequent studies adopted Centrarchidae as the family rank name and presented nested classifications that were meant to imply evolutionary relationships (Jordan 1877; McKay 1881; Bollman 1891). Jordan’s (1877) classification had two subfamilies with one containing Micropterus and all other genera were classified in the second subfamily, Lepominae. Within the Lepominae, the genera Ambloplites, Archoplites, Acantharchus, and Chaenobryttus (Lepomis gulosus) were grouped together. The remaining Lepomis species were classified into five genera that are no longer recognized, and Enneacanthus, Centrarchus, and Pomoxis were placed in the same grouping. The classification presented by McKay (1881) did not include taxonomic ranks above genus. Previous to this classification all Lepomis species were classified into eight genera, Chaenobryttus, Apomotis, Xenotis, Bryttus, Helioperca, Xystroplites, Eupomotis, and Lepomis. McKay (1881) placed all of these species, except for L. gulosus, into Lepomis. The classification presented by Bollman (1891) is important in that it recognized three subfamilies, Centrarchinae, Lepominae, and Micropterinae that are still in use (Table 1.1). Bollman’s (1891) proposed Centrarchinae contained Centrarchus and Pomoxis, whereas the Lepominae contained Archoplites, Ambloplites, Chaenobryttus, Acantharchus, Enneacanthus, Mesogonistius, and Lepomis. Lepomis gulosus was retained in Chaenobryttus as it was considered distantly related to other Lepomis species. The Micropterinae contained two recognized Micropterus species. After the studies of McKay (1881) and Bollman (1891), pharyngeal jaw morphology provided important information for hypotheses of relationships among Lepomis species. Hypertrophied lower pharyngeal arches were presented as evidence to remove L. gibbosus from Lepomis and into the genus Eupomotis (Richardson 1904). A later review of the pharyngeal arches resulted in an amplification of Bollman’s (1891) proposal that all species previously classified in Apomotis, Xenotis, Bryttus, Helioperca, Xystroplites, and Eupomotis were closely related and all species from these genera were placed in Lepomis (Bean and Weed 1911). However, as pointed out by Bailey (1938), some of the discussion in Richardson (1904) and Bean and Weed (1911) was based on hybrid individuals. The first representation of centrarchid relationships presented as a branching dendrogram was in Schlaikjer’s (1937) “suggestions on the phylogeny of the recent Centrarchidae.” This schematic of centrarchid relationships did not result from the current concept of a phylogenetic analysis, but was inferred from body depth, relative mouth size, and numbers of rays and spines on the dorsal and anal fins (Figure 1.18). In this “phylogeny” Centrarchus macropterus was depicted as the ancestral centrarchid species with Pomoxis and Archoplites being represented as early splits from this ancestral lineage. Elongated body shape was an important characteristic that motivated the grouping of Acantharchus, Ambloplites, and Micropterus (including the invalid Huro) (Figure 1.18). In contrast to the classifications of McKay (1881) and Bollman (1891), Schlaikjer (1937) classifies Lepomis species (except L. gulosus) into three genera (Lepomis, Apomotis, and Eupomotis) that were depicted as a group in the branching diagram (Figure 1.18). The Enneacanthus species, including
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Mesognistius chaetodon
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Chaenobryttus glosus
Enneacanthus gloriosus
Pomoxis sparoides VI
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Figure 1.18 Phylogeny of Centrarchidae presented in Schlaikjer (1937).
Pomoxis annularis
Eupomotis gibbosus
Lepomis auritus
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'Apomotis' symmetricus X 11-12
Schlaikjer(1937)
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Ambloplites rupestris
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Huro salmoides
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14
Centrarchid fishes
Icthelis
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Bailey (1938)
Helioperca Allotis
Bryttus
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Lepomis
Enneacanthus
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Lethogrammus Apomotis
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op litin i
an
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omi
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ra
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nt
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th
Acantharchus
Cent rar ch ina e
Lep om in
?
rini
pte
? Micro
ae
Archoplites interruptus
Figure 1.19 Phylogeny of Centrarchidae presented in Bailey (1938). Archoplites interruptus redrawn from Girard (1858).
the invalid Mesogonistius, were placed as closely related to Lepomis, with L. gulosus (in Chaenobryttus) outside of this group (Figure 1.18). Bailey (1938) in an unpublished Ph.D. dissertation presented a classification of Centrarchidae and a “hypothetical phylogeny” for the group that was depicted as a branching diagram (Figure 1.19). The characters used by Bailey (1938) were primarily anal fin spines, branchiostegal rays, dentition, body shape, opercle serration, scale morphology, and gill raker morphology. By classifying Centrarchus, Pomoxis, Archoplites, Acantharchus, and Ambloplites in the subfamily Centrarchinae, Bailey (1938) was the first author to propose that species in these genera are closely related (Figure 1.19). The genera Chaenobryttus (L. gulosus), Lepomis, Enneacanthus, Mesogonistius (Enneacanthus chaetodon), Micropterus, and Huro (M. salmoides) were classified together in the subfamily Lepominae (Figure 1.19). Given the uncertainty of Bailey’s (1938) placement of Enneacanthus and Mesogonistius in the phylogenetic tree (Figure 1.19), Lepomis and Chaenobryttus were depicted as the sister lineages and most closely related to Micropterus. Eight subgenera were proposed for ten recognized Lepomis species. Sister species pairs proposed within Lepomis were L. cyanellus–L. symmetricus, L. macrochirus–L. humilis, L. gibbosus–L. microlophus, and L. megalotis–L. marginatus. A detailed “phylogeny” of Lepomis species, as proposed by Bailey (1938), is given in Figure 1.20. A “theoretical phylogeny” was presented in a taxonomic revision of Micropterus that described four new species and subspecies (Figure 1.21; Hubbs and Bailey 1940). This Micropterus “phylogeny” was intuitively derived and based on character variation in scale row and fin counts, and pigmentation patterns (Figure 1.21). In this phylogeny Huro was still used as a monotypic genus to contain M. salmoides. Also, the subspecies of M. punctulatus were not presented as a group that is most closely related to one another (Figure 1.21). This is explained by the fact that Hubbs and Bailey (1940, p. 41)
Species diversity, phylogeny and phylogeography of Centrarchidae
Bailey (1938)
15
L. gulosus L. cyanellus L. symmetricus L. punctatus L. gibbosus L. microlophus L. humilis L. macrochirus L. auritus
Lepomis macrochirus
L. megalotis L. marginatus
Figure 1.20 Detailed phylogeny of Lepomis presented in Bailey (1938). Subgenera of Lepomis in Figure 1.19 were translated to species names using tables presented in Bailey (1938). Lepomis macrochirus redrawn from Forbes and Richardson (1920).
did not rule out a scenario where M. coosae originated through hybridization and introgression between M. dolomieu and M. punctulatus. This explains the “paraphyletic” depiction of M. punctulatus in their branching diagram (Figure 1.21). The phylogeny presented in Bailey (1938) was slightly modified and used to study the evolution of dorsal fin supports in percoid fishes, and particularly in Centrarchidae (Figure 1.22) (Smith and Bailey 1961). In this branching diagram Archoplites is resolved as the sister species of a group containing Pomoxis and Centrarchus (Figure 1.22). The phylogenetic position of Enneacanthus (Mesogonistius was no longer recognized) was still unresolved, but was hypothesized as closely related to Lepomis and Micropterus and not to genera in Centrarchinae (Smith and Bailey 1961). The lateralis system and osteology provided characters for inferences regarding centrarchid phylogeny (Branson and Moore 1962). In this study, relationships were proposed separately for centrarchid genera, species in Lepomis, and species in Micropterus. The “hypothetical dendography” presented as a phylogeny among centrarchid genera is quite different from the hypotheses presented in Bailey (1938) (Figure 1.19) and Smith and Bailey (1961) (Figure 1.22). In Branson and Moore’s (1962) hypothesis, Centrarchinae, as proposed by Bailey (1938), is paraphyletic relative to the Lepominae, and Chaenobryttus is nested outside of a sister group containing Lepomis and Micropterus (Figure 1.23a). Within Lepomis, Branson and Moore (1962) converge on a hypothesis of relationships that is less resolved than Bailey’s (1938) (Figure 1.20), but agree with Bailey (1938) in presenting the L. cyanellus–L. symmetricus and L. macrochirus–L. humilis species pairs (Figure 1.23b). The proposal of relationships among Micropterus species presented by Branson and Moore (1962) agrees with that of Hubbs and Bailey (1940) (Figure 1.21) in depicting M. salmoides as the sister species to all other Micropterus species. Also, Branson and Moore (1962) provide a phylogenetic hypothesis for M. treculi and M. notius (Figure 1.23c), two species that were either not recognized or not described when Hubbs and Bailey (1940) revised Micropterus. The last of the pre-cladistic hypotheses of centrarchid relationships discussed in this review was published after the development of cladistic methods, but is a verbal hypothesis of relationships among Micropterus species based primarily on pigmentation and ecological characteristics (Ramsey 1975). Three lineages in Micropterus were identified and Ramsey’s (1975) hypothesis was converted into a generally unresolved phylogeny. In this tree M. salmoides is grouped by itself, M. coosae and M. dolomieu are sister species, and M. punctulatus, M. treculi, M. notius, and M. cataractae are placed in an unresolved grouping (Figure 1.23d).
16
Centrarchid fishes
Hubbs and Bailey (1940)
Huro salmoides
M.d. dolomieu
M.d. velox
M. coosae
M.p. wichitae M.p. henshalli
M.p. punctulatus
Micropterus salmoides
Figure 1.21 Phylogeny of Micropterus presented in Hubbs and Bailey (1940). Micropterus salmoides redrawn from Forbes and Richardson (1920).
1.4.2 Phylogenetic hypotheses derived from analysis of character data The preceding section reviewed ideas about centrarchid evolutionary relationships that were intuitive, and did not utilize forms of character optimization seen in the current practice of phylogenetic systematics (Swofford et al . 1996). This section reviews more recent hypotheses of centrarchid relationships, and includes those that use a particular optimality criterion to analyze a coded character dataset. As a result of the publication of several studies and datasets over the past 30 years, Centrarchidae has come to provide an exciting system to investigate very relevant issues in systematics such as character congruence among independent molecular datasets (Near et al . 2004), the use of fossil data for calibrating molecular phylogenies (Near et al . 2005b), and the optimal use of phylogenies and divergence time estimates in comparative studies (Bolnick and Near 2005; Collar et al . 2005). The first studies of centrarchid relationships that used a defined optimality criterion to analyze a comparative data matrix were also the first studies to use genetic data in reconstructing centrarchid phylogeny. Allozymes, which are alternative forms of an enzyme produced by different alleles of a given locus that are usually detected by protein electrophoresis, were used to investigate relationships among centrarchid genera and among Lepomis species (Avise and Smith 1974b, 1977; Avise et al . 1977). In these studies allozyme variation was converted to pair-wise genetic distances, and the unweighted pair-group method (UPGMA) was used for cluster analyses that resulted in branching dendrograms. The phylogeny resulting from the UPGMA analysis that included the greatest taxon sampling among these studies is presented in Figure 1.24a (Avise and Smith 1977). These analyses agreed with earlier, pre-cladistic hypotheses by presenting Micropterus and Lepomis as sister lineages (Bailey 1938; Branson and Moore 1962), but Pomoxis grouped with this clade instead of with other
Species diversity, phylogeny and phylogeography of Centrarchidae
17
Smith and Bailey (1961) Pomoxis
Centrarchus
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IN A
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EN
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Micropterus Enneacanthus chaetodon
Figure 1.22 Phylogeny of Centrarchidae presented by Smith and Bailey (1961). Enneacanthus chaetodon redrawn from Smith (1907).
Centrarchinae (Acantharchus, Archoplites, Centrarchus, and Ambloplites). However, it is important to note that the branch length for this node in the allozyme genetic distance dendrogram was very short (Avise and Smith 1977, Figure 5). The allozyme phylogenies differ from several of the earlier morphological hypotheses in having Enneacanthus closely related to genera comprising Bailey’s (1938) concept of Centrarchinae (Acantharchus, Archoplites, Centrarchus, and Ambloplites), and not Lepomis and Micropterus. Also, relationships within Lepomis were different from the hypotheses presented in Bailey (1938) and Branson and Moore (1962) (Figures 1.20 and 1.23a, b), perhaps most notable is that L. gulosus was nested well within Lepomis, and not in a separate clade that would warrant recognition of Chaenobryttus. A later allozyme study that used a distance Wagner method to construct a centrarchid phylogeny (Parker et al . 1985) resulted in a fairly similar tree (Figure 1.24b). One noticeable difference was the nonmonophyly of Lepomis, a result that may have been an artifact of the genetic distance calculations or the distance clustering method used in this study (Figure 1.24b). Characters from kidney morphology, anal fin spine counts, and olfactory organ morphology were used in the first explicit cladistic analysis of centrarchid phylogeny (Mok 1981). Two separate trees were presented, as Mok (1981) did not combine all the morphological characters for one cladistic analysis. The first phylogeny lacked resolution and was based on five characters from kidney morphology (Figure 1.25a). The presence of an extreme posterior kidney was interpreted as a shared derived character (synapomorphy) for all Centrarchidae except Micropterus. The phylogeny has a basal polytomy with the outgroup taxon (Elassoma), Micropterus, and all other centrarchid genera (Figure 1.25a). Despite the lack of phylogenetic resolution Mok’s (1981) analysis of kidney morphology resulted in a clade containing Centrarchus and Pomoxis that agreed with earlier pre-cladistic hypotheses (Bailey 1938; Smith and Bailey 1961; Branson and Moore 1962). Mok (1981) stressed that the kidney morphology does not support the previous hypotheses that presented Lepomis and Micropterus
18
Centrarchid fishes
(a) Branson and Moore (1962)
(b) Branson and Moore (1962) L. cyanellus L. symmetricus
Archoplites Acantharchus Pomoxis
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M. dolomieu
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M. coosae M. punctulatus
M. coosae M. treculi M. punctulatus M. treculi
M. notius M. cataractae
Figure 1.23 (a) Phylogeny of centrarchid genera presented by Branson and Moore (1962). (b) Phylogeny of Lepomis presented by Branson and Moore (1962). (c) Phylogeny of Micropterus presented by Branson and Moore (1962). (d) Phylogeny of Micropterus converted from a verbal hypotheses presented by Ramsey (1975).
as sister taxa (Bailey 1938; Smith and Bailey 1961; Branson and Moore 1962). The second phylogeny in Mok (1981) was based on two characters, the number of anal spines and folding of the olfactory sac, as presented in Eaton (1956). This tree was also unresolved, but it did argue that more than three anal fin spines was a synapomorphy for Ambloplites, Acantharchus, Archoplites, Centrarchus, and Pomoxis (Figure 1.25b), a result that agreed closely with Bailey’s (1938) concept of Centrarchinae (Figures 1.19 and 1.25b). An undefined set of morphological characters was used for a cladistic analysis of centrarchids, and the resulting tree served as the basis for a comparative study of diet, functional feeding morphology, and behavior (Lauder 1986). The phylogeny had a basal polytomy with Micropterus, Lepomis, and clade containing Pomoxis, Centrarchus, Acantharchus, Archoplites, and Ambloplites (Figure 1.25c). Lepomis was monophyletic and L. gulosus was not closely related to Micropterus. In agreement with Bailey (1938), L. gibbosus and L. microlophus were sister species (Figures 1.20 and 1.25c) The next morphological phylogeny of Centrarchidae was presented in an unpublished Ph.D. dissertation and was based on cladistic analyses of 27 morphological characters (Chang 1988). This phylogeny was pectinate, or completely imbalanced, with Micropterus as the basal sister taxon to all other centrarchids (Figure 1.25d). One interesting aspect of this phylogeny was the placement of Enneacanthus as the sister taxon of the genera that comprise Bailey’s (1938) concept of Centrarchinae, and not closely related to Micropterus or Lepomis. Also, in agreement with several previous studies (Bailey 1938; Smith and Bailey 1961; Mok 1981; Parker et al . 1985), Centrarchus and Pomoxis were sister taxa (Figure 1.25d). Chang’s (1988) study is particularly important because it identified four morphological synapomorphies for Centrarchidae (exclusive of Elassoma), a posterior bifurcation of the swim bladder, the first hemal spine of the same length as the second, a deep groove on the first hemal spine, and contact between the first and second hemal spines.
Species diversity, phylogeny and phylogeography of Centrarchidae
(a) Avise and Smith (1977)
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(b) Parker et al. (1985) Acantharchus
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Figure 1.24 (a) Allozyme inferred phylogeny of Centrarchidae presented by Avise and Smith (1977). (b) Allozyme inferred phylogeny of Centrarchidae presented by Parker et al . (1985).
In a study examining the evolutionary patterns in functional morphological aspects of feeding in centrarchids, Wainwright and Lauder (1992) used a centrarchid phylogeny that resulted from a cladistic analysis of 53 undefined morphological characters. The tree was similar to that of Chang (1988) in that Micropterus is the basal sister taxon to all other Centrarchidae (Figure 1.26a). Also, in agreement with Bailey’s (1938) concept of Centrarchinae, this phylogeny presented Acantharchus, Ambloplites, Pomoxis, Centrarchus, and Archoplites as a monophyletic group (Figure 1.26a). Archoplites interruptus and C. macropterus were recovered as sister species, a relationship that had not been proposed in any of the previous hypotheses; however, Mok (1981) presented a tree based on olfactory organ folding that had a clade containing Pomoxis, Centrarchus, and Archoplites (Figure 1.25b). Interestingly, in Wainwright and Lauder’s (1992) phylogeny, Enneacanthus was nested within Lepomis and L. gulosus was the phylogenetically basal species in this clade. Some details of the relationships in Lepomis proposed by Wainwright and Lauder (1992) are consistent with previous hypotheses (Bailey 1938; Branson and Moore 1962), and others are unique to this analysis (Figure 1.26a). Mabee (1989, 1993) presented a phylogenetic analysis of Centrarchidae using 61 morphological characters. The trees were used to study the ontogenetic criterion in phylogenetics, asking if an ontogenetic series for a particular character provided a reasonable method to polarize the character in a phylogenetic analysis (Mabee 1989, 1993). From our own reanalysis of the data matrix and other published analyses of this dataset (Mabee 1993; Patterson 1996), it is clear that parsimony analysis using outgroup rooting results in hundreds (if not thousands) of most parsimonious trees. However, a single tree from the set of most parsimonious trees was selected for purposes of Mabee’s (1989, 1993) analyses of ontogenetic character evolution (Figure 1.26b). Despite the seemingly arbitrary nature of the selection of this tree, the strict consensus of the most parsimonious trees is quite well resolved (see Patterson 1996, Figure 1a), and is completely resolute with regard to the details of the phylogenetic relationships discussed in this review.
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Centrarchid fishes
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Figure 1.25 (a) Phylogeny of Centrarchidae based on a cladistic analysis of kidney morphology presented by Mok (1981). (b) Phylogeny of Centrarchidae based on a cladistic analysis of anal spine counts and scale morphology presented by Mok (1981). (c) Phylogeny of Centrarchidae based on a cladistic analysis of morphological characters presented by Lauder (1986). (d) Phylogeny of Centrarchidae based on a cladistic analysis of 27 morphological characters presented by Chang (1988).
The phylogeny presented by Mabee (1989, 1993) is interesting in many respects (Figure 1.26b). In agreement with two of the other morphological cladistic analyses (Chang 1988; Wainwright and Lauder 1992), Micropterus is the sister lineage of all other Centrarchidae. The relationships within Lepomis were very similar to that presented by Wainwright and Lauder (1992)—L. gulosus was the sister species to all others in the clade, and Enneacanthus was nested in Lepomis. Within Lepomis, Mabee’s (1989, 1993) phylogeny has two sister species pairs, L. megalotis–L. marginatus and L. microlophus–L. gibbosus seen in other phylogenies (Bailey 1938; Avise and Smith 1977; Lauder 1986; Wainwright and Lauder 1992); however, the sister species pairs L. macrochirus–L. humilis and L. cyanellus–L. symmetricus proposed by Bailey (1938) and Branson and Moore (1962) were not supported by these analyses (Figure 1.26b). In agreement with many of the earlier, pre-cladistic, morphological hypotheses (Bailey 1938; Branson and Moore 1962), a monophyletic Centrarchinae, exclusive of Enneacanthus, was present in the selected single tree from the pool of most parsimonious trees (Figure 1.26b). However, Acantharchus falls out of this clade in the strict consensus tree (Patterson 1996, Figure 1a). Over the past 5 years DNA data has increasingly been used in phylogenetic analyses of Centrarchidae. Three studies have focused on relationships of Micropterus species (Johnson et al . 2001; Kassler et al . 2002; Near et al . 2003) and have produced fairly congruent results; however, there are some unresolved issues with regard to species recognition in the clade that are illuminated by these molecular studies. Johnson et al . (2001) analyzed the phylogeny of Micropterus species using a maximum parsimony analysis of restriction enzyme digests of whole mtDNA genomes. The monophyly
Species diversity, phylogeny and phylogeography of Centrarchidae
(a) Wainwright and Lauder (1992) Micropterus Acantharchus Ambloplites Pomoxis Centrarchus Archoplites L. gulosus E. gloriosus E. chaetodon E. obsesus L. cyanellus L. symmetricus L. humilis L. auritus L. megalotis L. marginatus L. macrochirus L. punctatus L. microlophus L. gibbosus
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(b) Mabee (1993) M. salmoides M. dolomieu M. notius M. punctulatus M. treculi M. coosae Acantharchus P. annularis P. nigromaculatus Centrarchus Archoplites A. rupestris A. cavifrons A. ariommus A. constellatus L. gulosus L. cyanellus L. macrochirus L. symmetricus E. chaetodon E. obsesus E. gloriosus L. humilis L. auritus L. marginatus L. megalotis L. punctatus L. gibbosus L. microlophus
Figure 1.26 (a) Phylogeny of Centrarchidae based on a cladistic analysis of morphological characters presented by Wainwright and Lauder (1992). (b) Phylogeny of Centrarchidae based on a cladistic analysis of 61 morphological characters presented by Mabee (1989, 1993).
of Micropterus was not tested as only a single outgroup species was used, but the phylogeny depicts M. salmoides as the sister species to all other Micropterus (Figure 1.27a). Near et al . (2003) presented a maximum likelihood analysis of DNA sequences from two mtDNA genes, cytb and ND2 that were collected from 50 individuals sampled from 8 Micropterus species. This maximum likelihood phylogeny was similar to the tree presented by Johnson et al . (2001), but differs primarily where the root was placed. This difference was most likely a consequence of the use of a single outgroup taxon. Also, Johnson et al . (2001) did not provide support values for nodes in the phylogeny, and Near et al . (2003) presented a phylogeny that had most of the interspecific nodes supported with high bootstrap pseudoreplicate scores (Figure 1.27b). In Near et al .’s (2003) tree M. dolomieu and M. punctulatus were sister species, and this clade was sister to the remaining Micropterus species (Figure 1.27b). Differing from Johnson et al . (2001), Near et al . (2003) found M. treculi as the sister species of a clade containing M. salmoides and M. floridanus. There are two aspects of the Micropterus phylogeny presented by Near et al . (2003) that support the recognition of M. floridanus as a species distinct from M. salmoides: (i) the two species exhibit reciprocally monophyletic mtDNA haplotypes, and (ii) the intraspecific branch lengths are shorter than those subtending the interspecific node (Figure 1.27b). Kassler et al . (2002) utilized the sampling of cytb and ND2 mtDNA sequences from Near et al . (2003), but added more M. treculi specimens and included M. henshalli in the phylogenetic analyses. Also, Kassler et al . (2002) presented phylogenies that are derived from the analysis of 19 polymorphic allozyme loci. The mtDNA maximum likelihood phylogeny yielded two very surprising results. First, two distinct M. treculi mtDNA haplotypes were discovered. One of
22
Centrarchid fishes
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(b) Near et al. (2003) M. punctulatus
Lepomis macrochirus 100 99
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Micropterus notius Micropterus treculi Micropterus salmoides
M. dolomieu 96 100 M. punctulatus
Micropterus floridanus 0.005 substitutions/site
Figure 1.27 (a) Phylogeny of Micropterus based on a cladistic analysis of restriction digests of whole mtDNA genomes presented by Johnson et al . (2001). (b) Phylogeny of Micropterus based on a maximum likelihood analysis of mtDNA gene sequences presented in Near et al . (2003). Outgroup species are not shown. A scale bar for the expected number of substitutions is given in the lower right, and numbers at nodes are percent recovery in a bootstrap analysis.
these was resolved as the sister taxon of the clade containing M. floridanus and M. salmoides, and the other was closely related to M. punctulatus. Second, M. henshalli was nested within the haplotypes sampled for M. coosae (Figure 1.28a). These two patterns could be attributed to mtDNA introgression, a process known to occur in fishes (Avise 2001), but the allozyme inferred phylogeny offers some important clues to the unexpected results in the mtDNA phylogeny (Figure 1.28b). In an unpublished study, we have screened 100 M. treculi from three locations within the species’ limited native range and found 49 individuals with the haplotype that is sister to the M. floridanus–M. salmoides clade, and 51 individuals with the haplotype that is closely related to M. punctulatus (Figure 1.28a). There is no geographic pattern within M. treculi as the two haplotypes were found in equal frequency within the three populations sampled. However, regardless of which of the two divergent mtDNA haplotypes are found in a given M. treculi specimen, there is virtually no intraspecific variation among allozyme alleles or DNA sequences from nuclear genes. In the allozyme phylogeny constructed using a frequency parsimony method (Swofford and Berlocher 1987), M. treculi is closely related to M. punctulatus, a result that is expected from the previous classification of M. treculi as a subspecies of M. punctulatus (Hubbs and Bailey 1942). The presence of a divergent mtDNA haplotype that is closely related to the M. floridanus–M. salmoides clade that has no counterpart in the nuclear gene phylogeny (Figure 1.28b) cannot be explained by human introductions of M. punctulatus into the native range of M. treculi. As it stands, the mystery of the two divergent mtDNA haplotypes in the background of what appears to be a homogenous nuclear genome of M. treculi will have to be solved in future studies. The case of M. henshalli, or the Alabama Spotted Bass, is equally puzzling as the pattern revealed in M. treculi. The mtDNA maximum likelihood phylogeny resolves M. p. henshalli as distantly related to M. punctulatus and the haplotypes are nested within M. coosae (Figure 1.28a). Micropterus henshalli and M. coosae are sympatric throughout the Mobile Basin (Mettee et al . 1996; Boschung and Mayden 2004) and the similarity of the mtDNA haplotypes would indicate a
Species diversity, phylogeny and phylogeography of Centrarchidae
(a) Kassler et al. (2002) (mtDNA)
23
(b) Kassler et al. (2002) (allozymes)
M. floridanus
M. coosae
M. salmoides
M. treculi M. notius M. cataractae
M. dolomieu
M. dolomieu M. floridanus
M. punctulatus
M. cataractae
M. notius
M. treculi M. henshalli M. salmoides M. henshalli M. coosae M. punctulatus 0.5 changes
M. treculi
Figure 1.28 (a) Phylogeny of Micropterus based on a maximum likelihood analysis of mtDNA gene sequences presented by Kassler et al . (2002). Outgroup species are not shown. (b) Phylogeny of Micropterus based on a frequency parsimony analysis of allozyme alleles at 19 loci presented by Kassler et al . (2002).
recent introgression of mtDNA from M. coosae to M. henshalli. The allozyme phylogeny resolves M. henshalli as the sister taxon of a clade containing M. punctulatus and M. treculi with a very long branch separating M. henshalli from this clade (Figure 1.28b). In our own work we have collected M. coosae and M. henshalli in sympatry in the upper Coosa River system. These individuals have very similar mtDNA haplotypes (1.3% uncorrected genetic distance), but despite the sympatry of these species the M. p. henshalli haplotypes cluster together exclusive of the paraphyletic M. coosae mtDNA haplotypes (Figure 1.28b). Even if mtDNA introgression is obscuring the true phylogeny of M. henshalli, it is apparent that it is quite distinct from M. punctulatus at nuclear encoded loci (Figure 1.28b) and exhibits substantial morphological divergence (Gilbert 1973). Complete coding sequences from the mtDNA cytb gene were used to examine intraspecific and interspecific relationships of Lepomis species (Harris et al . 2005). All species in the clade, except the very recently elevated L. peltastes, were sampled and multiple individuals were included from each sampled species. As reported by Harris et al . (2005), mtDNA haplotypes from five Lepomis species were not reciprocally monophyletic (Figure 1.29). However, for reasons outlined in the following text, we have found it necessary to reanalyze the cytb data from Harris et al . (2005). The phylogeny presented in this review was obtained using a Bayesian analysis similar to that used by Near et al . (2005b), and we present the phylogeny as a phylogram (Figure 1.29). Harris et al . (2005) state that introgression and the presence of cryptic species best explain the pattern of extensive nonmonophyly observed in Lepomis species. Despite a reasonable probability for this scenario, hybridization cannot be detected without genetic data from nuclear genes or morphological analyses (Neff and Smith 1979; Shaw 2002), and identification of cryptic species would minimally require some degree of assessment of morphological divergence, but such data were not presented (Harris et al . 2005). One possible explanation for the nonmonophyly of Lepomis species not
24
Centrarchid fishes
Harris et al. (2005) * *
L. auritus group A
* * *
L. punctatus L. miniatus
* L. microlophus * L. miniatus
*
L. gulosus *
Illinois Oklahoma
*
*
L. symmetricus
L. cyanellus
* L. marginatus group A *
* * L. megalotis group A * L. marginatus group B * L. megalotis group C * Maryland L. auritus group B North Carolina * Maryland L. gibbosus West Virginia Minnesota
*
*
*
* L. m. macrochirus
* 0.05 changes
L. auritus group C L. m. mystacalis L. humilis
*
Figure 1.29 Phylogeny of Lepomis based on a reanalysis of mtDNA gene sequence data presented by Harris et al . (2005). The phylogram resulted from a Bayesian mixed model analysis. Outgroup species are not shown. A scale bar for the expected number of substitutions is given in the lower left, and asterisks at nodes indicate support with significant (0.95) Bayesian posterior probabilities.
explored by Harris et al . (2005) is misidentification of specimens. For example, the haplotype of L. symmetricus sampled from McCurtain Co., Oklahoma is very similar (low genetic divergence) to the haplotypes sampled from L. cyanellus (Figure 1.29). Given that L. cyanellus is sympatric with L. symmetricus in this region of Oklahoma (Miller and Robison 2004), specimens of L. cyanellus from the same location as this divergent L. symmetricus haplotype were not sampled, and Harris et al . (2005) do not state that they verified the identification of these specimens together, which means that specimen misidentification cannot be ruled out. The same circumstance can possibly be applied to the phylogenetic resolution of haplotypes from L. auritus group B that nests in the same clade as the sampled L. gibbosus haplotypes (Figure 1.29). The striking similarity of the haplotypes in L. auritus group B and L. gibbosus, in addition to the fact that the two species are sympatric where the L. auritus group B specimens were collected, points to a possible instance of specimen misidentification. Ancestral polymorphism was not considered as a mechanism that could result in the pattern of extensive species nonmonophyly observed in the Lepomis phylogeny (Figure 1.29). Ancestral polymorphism can result in nonmonophyly of a species’ alleles when the ancestral species is polymorphic at the locus, and the random sorting of the alleles during the splitting into multiple daughter species results in a gene tree that is incongruent with the species phylogeny (Neigel and Avise 1986; Pamilo and Nei 1988; Wu 1991; Hudson 1992; Hudson and Coyne 2002). The time to reach coalescence, when the species haplotypes are monophyletic, is proportional to the effective population size. Due to maternal inheritance, the coalescent time for mtDNA haplotypes is one quarter that expected for alleles of an autosomal locus (Moore 1995). One heuristic method to assess if ancestral polymorphism is driving a phylogenetic result is to determine if interspecific branches
Species diversity, phylogeny and phylogeography of Centrarchidae
25
(genetic distances) are longer than intraspecific branches in the phylogeny, with the assumption that long interspecific branches indicate that sufficient time has elapsed to expect coalescence and reciprocal monophyly (Moore 1995). The paraphyly of L. miniatus and L. marginatus–L. megalotis are in regions of the Lepomis phylogeny that have fairly short interspecific branch lengths relative to the intraspecific branch lengths, so ancestral polymorphism should not be ruled out as a cause for the observed paraphyly of these species. The first phylogenetic investigation among centrarchid genera using DNA sequences was an analysis of the mitochondrial cytb gene by Roe et al . (2002). The importance of this study was limited by the sampling of only one half of all centrarchid species, and by sparse phylogenetic resolution. Two recent studies have examined relationships of all extant centrarchid species, except the recently elevated L. peltastes and M. henshalli, using DNA sequences from multiple genes. Near et al . (2004) presented phylogenetic trees resulting from maximum parsimony and Bayesian analyses of a three gene data set consisting of the mtDNA, ND2, and two nuclear genes (S7 ribosomal protein intron 1 and the protein coding Tmo4C4 ). Two important conclusions were discussed in Near et al . (2004). First, separate analyses of each of the three sampled gene regions resulted in very similar phylogenies that indicated little incongruence between mtDNA and nuclear gene trees. Second, Shimodaira–Hasegawa tree topology tests indicated that 13 of 20 previous hypotheses of centrarchid relationships examined were significantly different from the best tree that resulted from the Bayesian analysis of the mitochondrial and nuclear gene dataset (Table 1.2). This allowed a unique perspective on how these earlier hypotheses compared in the context of a large set of characters that were sampled for most of the species level diversity in Centrarchidae. The phylogenies inferred from mitochondrial and nuclear gene DNA sequences demonstrated the monophyly of all polytypic genera, and in agreement with earlier studies resolved Lepomis and Micropterus as sister lineages (Bailey 1938; Smith and Bailey 1961; Branson and Moore 1962; Avise and Smith 1977), and provided strong support for a clade containing Enneacanthus Centrarchus, Archoplites, Ambloplites, and Pomoxis (Figure 1.30a, b). Other interesting relationships resolved in these analyses included Archoplites and Ambloplites as sister taxa, and the identification of two sister species pairs within Ambloplites. Relationships within Lepomis were highly resolved and most nodes received strong support in maximum parsimony bootstrap analysis or had significant Bayesian posterior probabilities (Figure 1.30a, b). The sister species pairs L. cyanellus–L. symmetricus and L. humilis–L. macrochirus, proposed by Bailey (1938) and Branson and Moore (1962) (Figures 1.20 and 1.23b), were strongly supported in the mtDNA and nuclear gene phylogenies (Figures 1.30a, b). Also, L. megalotis and L. marginatus were resolved as sister species, supporting the results from several earlier studies (Bailey 1938; Avise and Smith 1977; Mabee 1993). Previous investigations of Lepomis phylogeny have hypothesized that L. microlophus and L. gibbosus are sister species. This relationship was not supported in the mtDNA and nuclear gene phylogenies (Figure 1.30a, b). These two species are the only Lepomis species that exhibit specialized diets, feeding primarily on snails. Both L. microlophus and L. gibbosus possess morphological and behavioral specializations that function in crushing snails (Lauder 1983, 1986; Wainwright and Lauder 1992), and many of these characters had been used as evidence of common ancestry for these two species (Bailey 1938; Branson and Moore 1962; Lauder 1986; Wainwright and Lauder 1992; Mabee 1993). These phylogenies indicated that the evolution of these characters involved with molluscivory have a more complex evolutionary history than previously hypothesized. The dataset used in Near et al . (2004) was expanded to include one additional mitochondrial gene (16S ribosomal RNA) and two additional nuclear genes (calmodulin intron 4 and rhodopsin) for a total of 5553 base pairs of aligned DNA sequence data (Near et al . 2005b). The purpose of this study was to use fossil information to calibrate the molecular phylogeny to estimate divergence times in Centrarchidae. Ten centrarchid fossils were used to provide minimal age estimates for nodes in the phylogeny. Using a fossil cross-validation method (Near and Sanderson 2004; Near et al . 2005a), Near et al . (2005b) were able to identify four fossil calibrations that provided inconsistent molecular age estimates, and six consistent centrarchid fossils were used to calibrate the molecular phylogeny. Molecular divergence times of centrarchid species were estimated using penalized likelihood, a method that account for lineage specific molecular evolutionary rate heterogeneity (Sanderson 2002). The centrarchid phylogeny was presented as a chronogram, where the branch lengths are drawn to reflect estimates of absolute evolutionary ages (Figure 1.31). Given the temporal context of centrarchid diversification, Near et al . (2005b) point out that the origin of Centrarchidae at approximately 35 mya in the late Eocene–early Oligocene corresponds to a time of major global climate change to cooler conditions, and a signature in the fossil record of both lineage extinction and origination for many disparate clades across the tree of life. Another important result from the centrarchid chronogram that was exploited by later studies of functional character evolution and patterns of post-zygotic reproductive isolation was the finding that the major centrarchid lineages had
26
Centrarchid fishes
Table 1.2 Shimodaira–Hasegawa tests of alternative phylogenetic hypotheses of centrarchid fishes. Significant results are presented with an asterisk. Hypothesis
p
Three gene Bayesian phylogeny; Figure 1.30b
–
Schlaikjer (1937); Figure 1.18
30◦ C. Based on Diana’s results with largemouth bass, one might expect that other centrarchids would have similar tolerance for moderate diel fluctuations. Whitledge et al . (2002) measured routine metabolism for two size classes of smallmouth bass (50–100 g and 150–280 g) at 18, 22, 26, and 30◦ C. They found the general trends that others have found with respect to weight and temperature. Specifically, they revealed little difference between rates at 22 and 26◦ C for both size classes, an outcome observed in other studies that is somewhat contrary to the strict exponential relationship normally expected (Kitchell et al . 1977). Regression models developed for each temperature produced weight-dependent exponents (b values) from 0.53 to 0.86. Zweifel (2000) measured routine metabolic rate for white crappie ranging from 50 to 300 g at 18, 21, 24, 27, and 30◦ C. Bajer et al . (2004a) performed regression analysis on the Zweifel data to develop a metabolic rate model as a function of temperature and weight (b = 0.377; this value is much lower than the range of other studies of about 0.6–0.9). In more recent work, Bajer (2005) determined that the weight coefficient for white crappie to be a more reasonable 0.70. Woodley and Cech (personal communication with Christa Woodley, University of California at Davis) have been investigating metabolic rates and other physiological responses of Sacramento perch to high temperature and other environmental stressors (e.g., DO, pH, and salinity). Based on the habitat in which they historically resided, it is believed that Sacramento perch have a high tolerance for extreme temperatures. From laboratory and field observations they were able to determine optimal temperatures for various life stages (larvae, juvenile, and adults) and found significant ontogenetic differences. The exponential function has often been used to describe the relationship between fish metabolic rate and temperature (Figure 7.2), but other relationships have also been used. In recently published work, Gillooly et al . (2002) discuss how metabolic rate is expected to vary with temperature as well as body size. They use the Boltzmann factor, also known as
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the Van’t Hoff-Arrhenius relation, to express how rates of chemical reactions that combine to determine metabolic rate are expected to vary with changes in absolute temperature: M ∝ e−E/kT where M is metabolic rate, the symbol ∝ means “is proportional to,” E is the activation energy of the chemical reaction [about 0.63 eV (electron volts)], k is Boltzmann’s constant (8.6173 × 10−5 eV K−1 ), and T is absolute temperature (K). Gillooly et al . (2002) recommend the use of the Boltzmann factor to account for the effects of temperature on metabolic rate. They propose a form in which the temperature adjustment is expressed relative to that at some reference temperature. Over a typical range of biological temperatures (0–35◦ C), the effects of temperature as expressed in the Boltzmann factor are similar to those predicted by an exponential function of temperature (Figure 7.4). Brown et al . (2004) presented an outline of a metabolic theory of ecology. They showed how the effects of body size and temperature on metabolic rate could provide an explanation for a wide range of phenomena in ecology, from the individual level to the population and ecosystem levels. Brown et al . (2004) do not mention the substantial amount of work in fish bioenergetics that had come to the same conclusion about the importance of body size and temperature as major factors affecting metabolic rate (Kitchell et al . 1977 and subsequent work). West et al . (1997, 1999) provided an explanation for quarter-power allometric scaling of metabolic rate in animals, why metabolic rate per individual should be proportional to mass raised to the three-fourth power. They considered the transport of nutrients and other essential materials through fractal-like branching networks of tubes that service the entire body of an animal. Essential materials need to be exchanged with all cells in the body to keep them alive. They assume that natural selection has minimized the energy required to transport essential materials throughout the body, minimizing transport distances and times. They also assume that the size of the terminal tubes in the transport network is the same in all organisms (e.g., blood capillaries are the same size in mice and whales). They assume that natural selection has acted to maximize metabolic capacity. They reason that this leads to maximizing the total effective biological surface area through which essential materials are exchanged with all cells. The branching distribution network that creates this exchange area is volume-filling, leading to the consequence that “the area of the effective exchange surface scales as if it were a volume” (West et al . 1999, p. 1679). The fractal-like distribution network thus adds another dimension to living things, so that effective surface area scales as if it operated in three dimensions, and the corresponding biological volume scales as if it operated in four spatial dimensions. This is the reason that metabolic rate scales as mass raised to the three-fourth power, and why we should expect b = 0.75. For metabolic rate the exponent of mass is expected to be very close to b = 0.75 (West et al . 1997, 1999; Brown et al . 2004). Deviations far from this value should be regarded with suspicion. Large deviations may be due to statistical estimation problems caused by an insufficient range in body sizes. For example, in Figure 7.3, a large deviation from b = 0.75 is observed in the data from Shepard (1988), who used fish that differed in size by less than one order of magnitude. The deviation from b = 0.75 observed in the data from Wohlschlag and Juliano (1959) in Figure 7.3 is probably due to methodological issues. They measured wild-caught bluegill soon after capture, and if the stress and recovery from capture is size related, it could produce a stronger effect on the metabolic rate of larger fish and a deviation from b = 0.75. The theoretical basis provided by West et al . (1997, 1999) provides a strong justification, in our opinion, for using b = 0.75 as a standard value in fish bioenergetics models. This should also be the reference value against which to judge experimentally measured estimates.
7.4.2 Activity The effect of activity on metabolic rate has been investigated for a number of centrarchid species and is a significant factor in the development of an energy budget. The results of these studies can be used to derive activity multipliers or swimming dependent functions for respiration rate equations in bioenergetics models. Inclusion of an activity parameter in the standard metabolic rate equation is often accomplished in one of two ways. One way is by capturing the effect of activity as a simple multiplier of the routine (or standard) metabolic rate: M = aW b ecT ACT
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Centrarchid fishes
0 ln(I, Watts) Linear ln(I, Watts)
ln(R, Watts)
−1 −2 −3 −4
y = 0.0867x − 4.7797 R 2 = 0.999
−5 −6
0
5
10
15 20 25 Temperature (°C)
30
35
40
Figure 7.4 The effect of temperature on metabolic rate (M, in watts) as predicted by (i) the Boltzmann factor (also known as the Van’t Hoff–Arrhenius relation) and (ii) a simple exponential relation, M = a · exp(0.0867 · T), where a = exp(−4.7797) watts. The parameters of the exponential relation were estimated by linear regression of loge (M) on T.
where ACT is an activity multiplier usually between 1 and 2 (Hanson et al . 1997). Alternatively, active metabolism can be modeled as a function of swimming velocity: M = aW b ecT edV where V is swimming velocity (usually in either centimeter per second or body lengths per second) and d is a regression constant. Hanson et al . (1997) present an alternative formulation for incorporating swimming speed which is more complex, but not necessarily any more accurate. In their early studies on metabolic rates of bluegill, Wohlschlag and Juliano (1959) included level of activity as a factor in addition to temperature and fish size. The level of activity during the experiments varied from 0 to 24 cm s−1 . They derived mathematical models of metabolic rate by regressing log oxygen consumption on log body weight, swimming speed, and temperature. Data from Johnson and Charlton (1960) indicate that increased activity elevated respiration rate by a factor of about 1.5 for fingerling largemouth bass. They tested fish at predetermined temperature-specific maximum cruising speeds between 10 cm s−1 (5◦ C) and 40 cm s−1 (22◦ C). For tests at 5, 12, 17, 22, and 29◦ C, active metabolism exceeded standard metabolism by an average factor (i.e. ACT) of 1.54. Brett and Sutherland (1965) measured active metabolism of pumpkinseed sunfish in a tunnel respirometer to evaluate the relation between oxygen consumption and swimming speed. Tests were performed at several velocities between 3 and 37 cm s−1 . They found that for a 45-g fish at 20◦ C the active metabolic rate at the 60-min fatigue swimming speed of 37 cm s−1 was about nine times greater than the standard rate. Glass (1971) tested the effects of activity on four size classes of largemouth bass at a range of velocities ranging from a mean minimum of 7.4 cm s−1 to a mean maximum of 46.8 cm s−1 . The highest metabolic rate occurred at the higher velocities and exceeded that of fish at slower velocities by a factor of 4.2 for the smallest size class (30–40 g) and by a factor of 1.9 for the largest size class (88–95 g). Recent work by Bajer (2005) produced an updated metabolic rate equation for white crappie that includes weight, temperature, and activity as dependent variables in the form described earlier: M = 5.37 · W −0.296 e0.0606T e0.785V Cooke et al . (2001) tracked smallmouth bass with electromyogram transmitters that measured muscle activity to evaluate daily activity costs relative to the same estimates from mark-recapture and conventional telemetry studies. Their results indicate that smallmouth bass may travel nearly 100 times more than previously expected. They further concluded that based on bioenergetics modeling, this higher level of activity would require a 37% increase in daily food consumption to meet energetic requirements.
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Similar to increases in “voluntary” activity are energetic expenditures related to escape from predation. Cooke et al . (2003) investigated heightened stress in largemouth bass associated with fright-or-flight responses following simulated predation by avian predator models. They found elevated cardiac response, heart rate, and stroke volume of largemouth bass when presented with avian models and that this response was size-dependent. These nonlethal costs might need to be considered in bioenergetics models when predation avoidance is a significant factor. A recent review of studies using electromyogram telemetry lists 10 studies that included either smallmouth or largemouth bass (Cooke et al . 2004).
7.4.3 Hypoxia Metabolic rate is often affected by exposure to environmental stressors, such as low dissolved oxygen, turbidity, pH, and exposure to toxicants. One of the most common stressors experienced by warmwater fish and therefore one that is frequently studied is low dissolved oxygen or hypoxia. Cech et al . (1979) found a significant reduction in routine metabolic rate of adult largemouth bass as dissolved oxygen partial pressures declined. Critical oxygen tensions were achieved at progressively higher levels as temperatures increased from 20 to 30◦ C. To minimize largemouth bass mortalities in the wild, these results suggest that dissolved oxygen concentrations need to be maintained above 2.60 mg l−1 at 25◦ C and above 2.85 mg l−1 at 30◦ C. Moss and Scott (1961) exposed juvenile bluegill and largemouth bass to progressively lower dissolved oxygen concentrations while monitoring oxygen uptake rate to determine the critical dissolved oxygen level (i.e. concentration at which respiration rate decreased relative to a control) at 25, 30, and 35◦ C. They found critical levels increased from about 0.7 mg l−1 at 25◦ C to 1.0 mg l−1 at 35◦ C for bluegill and from about 0.8 mg l−1 at 25◦ C to 1.2 mg l−1 at 35◦ C for largemouth bass.
7.4.4 Turbidity Horkel and Pearson (1976) measured the effects of turbidity on ventilation rate and oxygen consumption of green sunfish at four temperatures (5, 15, 25, and 35◦ C) and several levels of turbidity. Ventilation rates eventually increased at all temperatures in response to elevated turbidity, with a response occurring at lower turbidities as temperature increased. However, oxygen consumption rates did not change over the range of turbidities tested at any of the four temperatures.
7.4.5 Environmental stressors Fish exposed to sublethal levels of environmental contaminants and other stressors often exhibit physiological and or biochemical responses. Shepard (1988) investigated the potential for using metabolic rate as an indicator of contaminant exposure in redbreast sunfish. Although fish exposed to sublethal levels of suspended contaminated sediment showed no difference in metabolic rate, those exposed to stream water with industrial effluent did have reduced metabolic rates. The author suggests this effect may have been a result of gill damage due to chlorine exposure and not a direct metabolic effect.
7.4.6 Schooling Just as there are conditions that result in suboptimal metabolic rates, there are also conditions that support a healthy metabolism. From the studies referred to earlier in this chapter one might generalize that optimal conditions for low metabolic rates include low temperatures, high dissolved oxygen, low turbidity, and minimal activity. There are other less obvious factors that affect metabolism. Parker (1973) investigated the relationship between metabolic rate and induced schooling for bluegill and largemouth bass (along with 13 other non-centrarchid species). He found that respiration
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rates for bluegill, but not for largemouth bass, were significantly lower when fish were grouped with conspecifics than when tested individually. These results correspond with what we know about the social behavior of these two species (i.e. bluegill commonly school and largemouth bass are more solitary). To the contrary, Wohlschlag and Juliano (1959) found grouped bluegill had a higher average metabolic rate than individuals in spring but no differences in summer. They suggest that this might be explained by relaxation of competitive spawning interactions.
7.5 Energetic wastes (egestion, excretion, and SDA) Egestion, excretion, and specific dynamic action (SDA) are energetic costs associated with digestion and processing of consumed food. Egestion represents the loss of energy in that portion of consumed food that is not digestible, passes through the gut without absorption, and is defecated. Excretion represents energy loss in the form of ammonia/ammonium and urea when proteins are broken down for energy production or storage. SDA is the common name given to the increase in metabolic rate that is associated with digestion and processing of a recently consumed meal. Gerking (1971 and earlier works) and Savitz (1971) performed many of the early experiments on nitrogen excretion in centrarchid fishes. They investigated the relationships between nitrogen excretion and fish size and ration for bluegill. Beamish (1972) estimated both excretion and egestion rates for largemouth bass as a percentage of total energy consumed. Although it has been demonstrated that egestion and excretion are both functions of temperature and ration level (Elliott 1976), the sum of the two remains fairly constant (Kitchell et al . 1977). Therefore, in lieu of detailed data, both are usually estimated as a set proportion of consumed energy in most bioenergetics models. Rice et al . (1983) and Adams et al . (1982) derived their estimates of egestion (10.4% of consumed food) and excretion (7.9% of consumed food) from empirical studies by Beamish (1972). For a bluegill bioenergetics model, Breck (1993) adopted nonspecific values from Brett and Groves (1979) of 0.168 and 0.0841 as the proportion of ingested energy accounted for by egestion and excretion, respectively. Hanson et al . (1997) used values of 10.4% for egestion and 6.8% for excretion for adult largemouth and smallmouth bass. Although rare in bioenergetics models, Breck and Kitchell (1979) and Hanson et al . (1997) calculated both egestion and excretion as functions of temperature and ration for bluegill. In bioenergetics models, SDA is typically represented as a constant proportion of the energy consumed, although Beamish and Trippel (1990) argue that many studies show that this is not the case and doing so may result in inaccurate model outputs. Similar studies by Beamish (1974) and Tandler and Beamish (1981) reported slightly different mean apparent SDAs for largemouth bass, 14.2 and 11.3% of energy ingested, respectively. Both studies found the typical temporal pattern of a relatively rapid rise in oxygen consumption after feeding, reaching a maximum around 2 hours postfeeding and then a gradual decline back to baseline that may take 1 to 3 days. The period of elevated metabolic rate depended on fish size and meal size, but the energy demand relative to energy ingested was mostly unaffected by meal size or fish size. Pierce and Wissing (1974) found a mean energy cost of food utilization for bluegill of 12.7% when they were fed mayfly nymphs (Hexagenia limbata). Schalles and Wissing (1976) in a similar study found an SDA value for bluegill when fed a dry pellet diet to be about 14.9% of consumed energy. The original bluegill bioenergetics model of Kitchell et al . (1974) adopted the 13% value from Pierce and Wissing. In the original largemouth bass models, Rice (1981) and Adams et al . (1982) both used the value of 14.2% derived by Beamish (1974). The SDA values used in the latest version of the Fish Bioenergetics (3.0) software (0.172 for bluegill, 0.163 for largemouth bass, and 0.16 for smallmouth bass; Hanson et al . 1997) are not consistent with the earlier studies and their origin is unclear.
7.6 Growth energetics 7.6.1 Energy density and body composition Fish energy density can have a large effect on growth rate. Suppose two fish each have 2 kcal of energy (consumption minus losses) available for growth. If one fish has an energy density of 1 kcal g−1 then it can add 2 g of weight. If the
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other fish has an energy density of 2 kcal g−1 then it can add only 1 g of weight and keep the same energy density. Fish energy density was not explicitly included as a parameter in the sensitivity analyses of Kitchell et al . (1977) or Bartell et al . (1986), but it can have a large influence on predicted growth (Hartman and Brandt 1995). Bioenergetics models often do their accounting in terms of energy, keeping track of energy input, energy losses, and change in total body energy content, considering energy density of the food and the fish. When considering changes in energy density or the corresponding changes in proximate body composition, the issue arises of conversion from one chemical type to another. In particular, protein could be converted to lipid following deamination and further conversion, but lipid cannot be converted into protein without addition of nitrogen. This sets some limitations on the conversion of one chemical class into the other. But it is generally the case that fish growth is limited by available energy, not a shortage of nitrogen or phosphorus (Schindler and Eby 1997). Fish energy density depends on body composition, which is related to percent dry weight and proximate composition. Brett and Groves (1979) summarize much literature about fish energetics, including the energy values of body components such as lipid, protein, and carbohydrate. For mammals, the familiar value for heat of combustion of lipid is 9.45 kcal g−1 (or 39.55 kJ g−1 , where 1 kcal = 4.1855 kJ). Brett and Groves note that for fish, which have lipids that are more unsaturated than mammals, the corresponding value for lipid is lower: 8.66 kcal g−1 (36.25 kJ g−1 ). The heat of combustion of protein is considered the same for mammals and fish: 5.65 kcal g−1 (23.65 kJ g−1 ). When body protein is used as an energy source, as during starvation, the metabolizable energy is only 4.80 kcal g−1 (20.1 kJ g−1 ) because some energy is lost with the nitrogen excreted as ammonia. Brett and Groves give a value of 4.10 kcal g−1 (17.16 kJ g−1 ) for mammal and fish carbohydrate. However, the carbohydrate content of fish is usually much less than 1% of wet weight, so this component is often omitted in analysis of fish energy density. Recent work is showing differences in fatty acid composition among species and among sizes of fish within species, probably related to differences in diets (e.g., Iverson et al . 2002), and this would contribute to variation in the mean energy value of lipid. Fish total energy content (E, kJ) can be calculated from the energy values of the component tissues. Let F represent the gram of lipid (fat) and P represent the gram of protein in a fish. The total body energy will be given by the following equation, where Df is the energy density of lipid (kJ g−1 lipid) and Dp is the energy density of protein (kJ g−1 protein). E = F Df + P Dp Fish energy density (d = E/W , kJ g−1 ) can be calculated by dividing E by total wet weight (W , g). Let f = F /W represent lipid content as a fraction of total weight, and p = P /W represent protein content as a fraction of total weight. d = f Df + pDp From these equations it is clear that fish energy density depends on proximate composition. Energy density is expected to be linearly related to the fraction lipid, and therefore, energy density is expected to change with body condition. Fish in better condition generally have higher levels of lipid, which translates into higher energy density. Fish percent dry weight changes with ontogeny. Larval fish have a much higher percentage of water and a lower energy density compared to older and larger fish. Breck (1993) reported that bluegill smaller than 6 mm were about 10% dry weight, and the median value for fish from 8 to 31 mm was 18% dry weight. McComish (1974) measured percent dry weight in 100 bluegill ranging from 35 to 192 mm and observed a range from about 24 to 33% dry weight. When bluegill data for small individuals (4.4–31.2 mm TL; Breck unpublished) are combined with the data of McComish (1971, 1974) for larger individuals (37–192 mm TL), one can observe a clear trend for percent dry weight to increase with body length (Figure 7.5). Typical body composition and energy density of bluegill in relation to fish length can be estimated from the regression equations shown in Figure 7.5 and additional data and information from McComish (1971, 1974). Fraction water (h) was estimated from the regression equations shown in Figure 7.5 for wet weight (W ) and dry weight (Wd ) in relation to length (L). h = 1 − Wd /W log10 (Wd /W ) = −1.060 − 0.255 log10 (L). Analysis of McComish’s (1971) data for grams of water (H = hW ) and grams of protein (P ) revealed a very strong relationship: loge (P ) = aP + bP loge (H )
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1000 100 10 Wet, McComish Weight (g)
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Length (mm) Figure 7.5 Wet weight and dry weight of bluegill over most of the common length range for this species. Data for individuals from 37 to 192 mm (N = 100) are from McComish (1971); data for individuals from 4.4 to 31.2 mm (N = 141) are from Breck (unpublished). Regressions use the equation: log10 (W ) = a + b log10 (L), N = 241. For wet weight, a = −5.416 (95% CL: ±0.026), b = 3.331 (±0.016), R2 = 0.999; for dry weight, a = −6.476 (±0.034), b = 3.586 (±0.021), R2 = 0.998. For reference, the equation for bluegill standard weight is shown as a dotted line (a = −5.374, b = 3.316). Because the slope for dry weight (3.586) is larger than the slope for wet weight (3.331), longer fish tend to have higher fraction dry weight and lower fraction water than smaller fish.
where aP = −1.535 ± 0.016, bP = 1.040 ± 0.006; N = 100, R 2 = 0.999, so water content can be used to estimate protein content. McComish (1974) reported a very strong relationship between grams of ash (A) and length: loge (A) = aA + bA loge (L) where aA = −16.674, bA = 3.540; N = 100; R 2 = 0.99. Therefore, length can be used to estimate wet weight and dry weight (Figure 7.5), fraction water and grams of water, and grams of ash. Water content (H ) can be used to estimate protein (P ). Grams of lipid (F ) can be estimated by subtracting water, protein, and ash from wet weight. Proximate composition can then be expressed as fractions of body weight and plotted in relation to fish length (Figure 7.6). The fractions of protein and lipid can be used to calculate energy density, which decreases as fraction water increases (Figure 7.7). These figures show the general pattern of body composition as bluegill increase in length. Individuals that are heavier at a given length would be expected to have a smaller fraction water and a larger fraction protein and lipid. Fish percent dry weight typically changes with body condition. Anderson (1959) and McComish (1974) both found a significant positive correlation with condition factor (K = 105 · W/L3 ) in bluegill. Fish energy density can have a strong seasonal component, often highest in the fall and lowest at the end of winter. This has been noted in largemouth bass (Brown and Murphy 1995) and white crappie (Bunnell and Marschall 2003). The effect of such seasonal changes on bioenergetics model predictions depends on the question being asked. Stewart and Binkowski (1986) used their bioenergetics model of alewife to compare simulations that included seasonal changes in alewife energy density to simulations using a constant energy density. They found that estimates of annual production and consumption differed by less than 10%, but there were larger differences in the seasonal estimates of production and consumption. The studies of centrarchid life history by Bunnell and Marschall (2003) and Garvey and Marschall (2003) suggest that a seasonal pattern of energy storage can contribute to fitness.
Fraction water, protein, lipid, ash
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1.0 0.9 0.8 0.7 0.6 0.5 0.4 0.3 0.2 0.1 0.0 0
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Figure 7.6 Body composition of bluegill in relation to fish length, estimated from data of McComish (1971).
There are two common observations regarding energy density and body composition. The first is that percent lipid declines as percent water increases; the second and related observation is that energy density (J g−1 wet weight) declines linearly as percent water increases (Brett et al . 1969; Henderson and Ward 1978; Rottiers and Tucker 1978; Love 1980; Weatherley and Gill 1983; Van Pelt et al . 1997). Breck (1998, 2008) proposed an explanation for these observations based on a mass-balance constraint and the hypothesis that a certain amount of water is associated with each gram of protein, and another smaller amount is associated with each gram of lipid in the body. If these amounts of water per gram protein and water per gram lipid are constant (e.g., for a specific size of fish and outside the range of starvation), then it can be shown that percent lipid will vary linearly with percent water, and that energy density will be a linear function of percent water. Because percent water changes with body size (Figure 7.6), these expected linear relationships for lipid and energy
10.0
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Figure 7.7 Body composition and calculated energy density of bluegill in relation to fraction water, estimated from the data of McComish (1971). Energy density calculations assume 36.25 kJ g−1 lipid and 23.65 kJ g−1 protein.
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density can appear noisy or even curvilinear if data from a wide size range of fish are combined; including log(W ) in the analysis can reduce the unexplained variation (Breck 2008). Pearse (1924) was apparently the first to measure the body composition of entire fishes. Earlier workers had measured edible portions or other parts. Working at the University of Wisconsin, Pearse also seems to have been the first to measure the body composition of centrarchids (largemouth bass and pumpkinseed), which he captured in Lake Mendota. His sample sizes were very small. He fed two juveniles a variety of invertebrates and other foods for 63 days and compared them with one juvenile that was starved for 63 days and another that was just caught from the lake. The two fed fish approximately doubled their weight (initial mass: 15.0 and 27.1 g; final mass: 36.94 and 62.84 g), had a relatively modest amount of water (73.72 and 71.31%), a relatively modest amount of fat (2.88 and 6.53% wet mass), and a modest amount of ash (4.13 and 4.11%), compared to the fish that starved (from 60 to 46.3 g), which had 75.73% water, 7.24% ash, and 0.51% fat. The fish caught fresh from the lake on August 20 was 25.71 g, 77.11% water, 3.78% ash, and 1.46% fat. He also compared two starved pumpkinseed with one fed and one caught fresh in the lake. The one starved fish decreased from 75.3 to 55.82 g in 62 days, with a final 76.01% water, 6.93% ash, and 0.42% fat; the other starved fish decreased from 70 to 37.89 g in 72 days, with a final 73.39% water, 9.53% ash, and 0.44% fat (this fish was dead when measured, but was alive the previous day). The fish fed for 84 days had a final mass of 26.37 g, had 72.75% water, 4.39% ash, and 5.64% fat. The fresh-caught fish weighed 52.68 g, had 74.44% water, 4.96% ash, and 2.35% fat. Note that the increased percentage of ash in the starved fish was almost certainly due to a reduced amount of other material with a constant mass of ash, not an increase in the mass of ash. These data are consistent with more recent information (Gerking 1955; Lee 1966; Savitz 1971; Niimi 1972; Garvey et al . 1998; McCollum et al . 2003). Together they suggest that centrarchids have a relatively low amount of fat and a high amount of ash compared to salmonids (e.g., Rottiers and Tucker 1978). Gerking (1955) studied the influence of consumption rate on body composition and protein metabolism of bluegill. Bluegill fed mealworms at higher rates grew at higher rates and developed higher levels of lipid. Savitz (1971) studied nitrogen excretion and protein consumption by bluegill. Niimi (1972) described the changes in body composition of largemouth bass during starvation, and Savitz (1971) did the same for bluegill. They both found that lipid level decreased with starvation. Niimi (1974) described the relationship between ash content and body weight in largemouth bass (and two other species). McComish (1974) developed regression models to predict the proximate body composition of bluegills. Consumption rate’s influence on body composition and energy density probably provides part of the explanation for the consumption-dependent errors recently identified in bioenergetics models (see Sections 7.2.4 and 7.3.5). Stock fish placed on reduced rations would be expected to reduce their energy density over several days; a bioenergetics model that used a higher value of energy density appropriate for the initial stock fish would tend to underestimate growth (or overestimate weight loss). Similarly, stock fish placed on increased rations would be expected to increase their energy density and lipid level over several days; a model that used the (lower) level of energy density of the stock fish would tend to overestimate growth. These are the directions of model errors identified by Bajer et al . (2004b). Careful measurements of energy density before and after such experiments would indicate how much of the model error could be explained by such expected dynamic changes in energy density. Hartman and Brandt (1995) describe methods of estimating energy density of fish. One recent method uses inductance, which varies with the proportion of water in the body, to estimate body composition. Lipid content has a large influence on fish energy density. The measured value for lipid content can depend significantly on the chemical method used to extract the lipid (Randall et al . 1991). Standard methods are important. Few energetics models have tracked body composition in addition to weight. An early model that incorporated body composition was developed for the African catfish Clarias gariepinus (Machiels and Henken 1986, 1987). Garvey and Marschall (2003) modeled seasonal energy allocation to growth, fat, and gonad for largemouth bass populations at low and high latitudes, and Bunnell and Marschall (2003) developed a similar model for white crappie. Schindler and Eby (1997) considered the stoichiometry of fishes and their food and asked whether fish growth was likely to be limited by nitrogen or phosphorus. They note that in the unusually rapid growth in some aquaculture situations, fish growth can be limited by the availability of N or P in the food. Their analysis of 28 populations and 18 species revealed that growth was almost always limited by energy and not by N. In only 3 of 186 cases examined (1.6%) was growth limited by P.
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7.6.2 Maximum sizes In contrast to birds and mammals, fish have plastic or indeterminate growth. Growth continues as long as energy intake exceeds energy expenditure. For some fish species, there is a limit to the number and size of muscle fibers in the body (Weatherly and Gill 1987). For these species, growth in size can continue until this limit is reached. There are an increasing number of suggestions that the optimum growth rate for fish may be less than the maximum possible growth rate (see Section 7.3.3). If these suggestions turn out to be correct, then it may be rare that fish approach a maximum size for the species. Under the Von Bertalanffy model, all fish approach a maximum specified size for that population.
7.6.3 Condition indices The related topics of fish condition and growth in length are areas where recent progress has been made and where further development and application of bioenergetics models would be helpful. Fish condition refers to relative plumpness, fish weight in relation to length, hence simulating changes in fish condition require simulating changes in fish length as well as weight. Proximate body composition and energy density are also involved. Fish in good condition generally have higher levels of lipid, higher energy density, and are more resistant to starvation than fish in poor condition. Female bluegills in poorer condition have lower fecundity than females in good condition (Breck 1996). Recent work on white crappie demonstrates that condition influences the rate of growth in length (Bajer 2005; Bajer and Hayward 2006). For a given increase in weight, fish in poor condition allocate more of the new tissue to improving condition and consequently grow less in length than fish in good condition (Bajer and Hayward 2006). Fish condition thus affects survival, reproduction and growth in length. Many different condition indices have been used on centrarchid fishes and here we briefly summarize some of those indices as well as centrarchid bioenergetics models involving changes in condition. Heidinger and Crawford (1977) showed that the liver-somatic index (liver weight as a percentage of wet weight) of largemouth bass varied with temperature and feeding rate. The relative size of the liver was shown to be an index of the average feeding rate over the past several days. Wege and Anderson (1978) introduced relative weight (Wr ) as a new index of condition. Standard weight (Ws ) reflects a reference weight for fish of a particular length (L) (Wege and Anderson 1978): Ws = aLb where a and b are constants for a given species. Relative weight (Wr ) expresses fish weight (W ) as a proportion (or percentage) of standard weight, and is an indication of the relative plumpness or condition of an individual compared to the standard. W Wr = Ws Murphy et al . (1991) proposed standard weight equations for several game fishes, including several centrarchids. Smith et al . (2005) used the changes in body shape that occur with starvation to devise a morphological index of nutritional status for larval largemouth bass (8.0–15.4 mm SL). They developed a multivariate index based on 23 characters that was able to correctly classify 92% of fed and 78% of unfed individuals. Most of the differences between fed and unfed fish occurred along the vertical body dimension, especially near the abdomen. A much simpler index, based on the ratio of body depth at the anus to standard length, was almost as effective, correctly classifying 83% of fed and 79% of unfed fish for the 7 days of the experiment. As expected, the magnitude of difference between fed and unfed fish increased with days of starvation. Weber et al . (2003) discuss several biochemical indices of condition in juvenile fishes. They discuss whole-body total lipids, whole-body triglycerides, muscle RNA:DNA ratio, and muscle protein. The novel aspect of their study is that all indices were measured for each individual fish. This permits analysis of how the suite of indices changes over time, for example, in comparing fed versus starved individuals. They give recommendations of methods for each biochemical
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index. For a starved group of juvenile rainbow trout, the RNA:DNA ratio decreased after just a few days, and whole-body lipids and triglycerides decreased to very low levels by day 18; however, mortalities did not begin until day 32 of their experiment. One early application of a bioenergetics model (Figure 7.1) was to a largemouth bass population in a power-plant cooling pond, where the model was used to better understand the factors that might be responsible for the poor condition (extreme thinness) of the bass at certain times of the year (Rice et al . 1983). Van Winkle et al . (1997) developed a model that included dynamic changes in body condition. Bajer (2005) and Bajer and Hayward (2006) showed very nicely that, for a given increase in weight, growth in length depends on relative condition. For the same level of consumption, fish in good condition had a greater rate of growth in length than fish in poor condition. They presented a method for estimating growth in both length and weight in white crappie using a modified bioenergetics model.
7.6.4 Stunting Stunting occurs in many centrarchid species, notably bluegill, black crappie, and largemouth bass. Stunted fish are small for their age, and their growth rates are low relative to some standard. Not all populations of small fish are stunted; if mortality rate is high, a population may have predominantly small fish despite rapid growth. The following comments are directed toward the issue of stunting, not small size per se. Several hypotheses have been advanced to explain stunting in centrarchids, including genetics, food availability per fish, and social influence of size at maturity of male bluegill. The energetics of growth are involved in several but not all of these hypotheses. The proximate cause of stunting is a low amount of energy used for growth. The more ultimate cause of such low energy is selection for life history strategies that maximize contribution of offspring to the next generation. This can result from high recruitment, without sufficient mortality to produce a good-growing density. One hypothesis is that genetic differences cause some populations to grow at a slower rate. This can definitely occur, as shown in common garden experiments involving young striped bass (Morone saxatilis) from various locations reared in identical conditions (Conover et al . 1997). However, in most cases of stunting in centrarchid populations, genetics seems unlikely to be the major cause of slow growth (e.g., pumpkinseed; Heath and Roff 1987). Similarly, the genetic hypothesis for stunting has been studied—and rejected—in fish populations from other fish families (e.g., yellow perch; Heath and Roff 1987). The major cause of stunting in centrarchid populations is a high density of juveniles, resulting in a reduced availability of food per individual. Evidence for this comes from observations and experiments. Fish increase their growth rate when moved from stunted populations to conditions with more food (Bennett et al . 1940, cited in Parker 1958). When the density of stunted populations is sufficiently reduced, either by winterkill (Beckman 1950) or experimental or management action (Parker 1958; Schneider and Lockwood 2002), the growth rate increases significantly. The effects of such actions or events typically last only a few years; growth rate slows over time, and the size structure returns to the stunted condition as the population biomass of fish increases (Schneider and Lockwood 2002). Similar responses occur in other fishes (e.g., roach Rutilus rutilus, Burrough and Kennedy 1979; bream Abramis brama, Wright 1990; white sucker Catostomus commersoni , Brodeur et al . 2001; Arctic charr Salvelinus alpinus, Amundsen et al . 2007). The growth rate of male bluegills depends in part on the size structure of the population. In bluegill, where there is strong female preference to mate with larger males, the sizes of males that are successful at mating depend on the size structure of the male population (see Chapter 5). If large male bluegills are sufficiently abundant in the nesting colony, then smaller males will be unsuccessful at mating (Jennings et al . 1997), so it is to their advantage to allocate more energy toward growth rather than reproduction until they attain a larger size. If large male bluegills are sufficiently rare or absent, then there may be a decline in male size at age as more male fish allocate a larger proportion of available energy to reproduction instead of growth. An experiment is currently underway in Illinois to examine the social hypothesis for fish maturing at a small size (see Chapter 5). Growth rate of juvenile centrarchids in the littoral zone can be independent of the growth rate of adults (Osenberg et al . 1992; Mittelbach and Osenberg 1993). Mittelbach and Chesson (1987) presented a two-stage population model for bluegills.
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7.6.5 Overwinter survival Growth can influence overwinter survival of centrarchid fishes in several ways. Growth during summer and fall influences fish body size as winter begins, and therefore affects the vulnerability to size-dependent predation. There also appears to be size dependence in the ability of some species to tolerate low temperature (Oliver et al . 1979). Growth influences the amount of energy reserves available to sustain the fish until food availability increases in the spring. Because of the allometry of respiration rate, larger fish use energy at a lower weight-specific rate than do smaller fish. In long-term controlled experiments that simulated exposure to typical winter temperature regimes, Oliver et al . (1979) found that young-of-year (YOY) smallmouth bass overwinter survival was directly related to size and body composition. Long fish survived better than short fish, and the ratios of mean dry weight to wet weight and ignitable weight to dry weight appear to be useful for differentiating between survivors and those that succumb to prolonged cold exposure. The standard fisheries approach to modeling survival is to account for natural mortality and fishing mortality, usually with constant age-dependent instantaneous rates (Ricker 1975). Although this is a useful approach for fisheries assessment, it can need modification when applied to bioenergetics models that focus on dynamic individual and population responses within a year. For example, bioenergetics models that account for changes in condition, or energy reserves permit the modeling of starvation mortality as an increased probability of death as condition declines or energy reserves are exhausted (Rice et al . 1983; Breck 1993, 1998; Kooijman 2000). Wright et al . (1999) concluded that the bioenergetics models they tested made poorer predictions for winter conditions than for summer. Adams et al . (1982) used a bioenergetics model of adult largemouth bass to evaluate the dynamics of food consumption, somatic and gonadal growth, energy storage, and activity throughout the year with emphasis on the overwinter period when prey abundance is lowest. They found that in a Tennessee reservoir during the winter, consumption and standard metabolism were low and no growth or energy storage occurred. During these periods of low food consumption, lipids were probably utilized from both the body and the viscera. Their modeling suggested that winter energy demands were greater than that which could be accounted for by catabolism of body tissue alone, and thus some consumption occurred throughout the winter at reservoir temperatures of 6 to 10◦ C. Garvey et al . (1998) used a combination of reservoir surveys of natural responses and experimental manipulation in reservoirs, ponds, and artificial pools to assess size-related overwinter survival of largemouth bass. They determined that the lower survival rate of the smaller individuals was more likely due to greater susceptibility to predation than for energetic reasons such as a lower amount of stored energy reserves. From field and laboratory observations with age-0 white crappie, McCollum et al . (2003) found that winter severity was a better indicator of overwinter survival than size, feeding level, or energy depletion. In laboratory studies, only 47% of all white crappies survived a simulated severe winter, whereas 97% survived a simulated mild winter. In severe winter, neither of the indicators listed earlier influenced mortality. They suggest that osmoregulatory failure as a result of exposure to temperatures colder than 4◦ C for at least 1 week may be responsible for high mortality rates.
7.7 Reproductive energetics There are several aspects of reproduction that have important energetic ramifications, including energetic costs of gonad development, nest building, and parental care. Most bioenergetics models do not account for somatic and gonadal growth separately, but typically include both in a single compartment. Increased activity costs due to spawning, and parental care also are rarely included in bioenergetics models.
7.7.1 Gonad development costs A bioenergetics model for centrarchids needs to account for the energy costs of reproduction, including gonad development and parental care. As in all fishes, reproductive costs of gonad development are much greater for females than for males. To the contrary, reproductive costs of parental care for centrarchids are much greater for males, because it is the males
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that make the nest, guard the eggs and larvae, and, in black bass, guard the fry for several weeks after they leave the nest. In bluegill, a variable proportion of young males become mature precocially (sneakers), allocating some energy to reproduction instead of growth, and probably foregoing some foraging during reproductive bouts. Species that spawn in early or late spring (e.g., largemouth bass, smallmouth bass, black crappie, and white crappie) tend to begin accumulating energy for reproduction in autumn, whereas species that spawn in summer (e.g., bluegill) can wait until spring to allocate energy to reproduction (James 1946; Morgan 1951a, 1951b; Adams et al . 1982; Bunnell and Marschall 2003; Garvey and Marschall 2003). Morgan (1951a, 1951b) measured the gonadosomatic index (GSI, gonad mass as a percentage of body mass) for bluegill, black crappie, and white crappie in Buckeye Lake, Ohio, from mid-March to late August, usually at intervals of 3 to 7 days (note: actually, he reported the ratio of body mass to ovary mass, which is the inverse of GSI—here, we have expressed his results in terms of the more familiar GSI). He observed that female bluegill GSI was very low (average GSI about 1.1–1.7%) during March and April, increased rapidly to about 7.7% between May 5 and May 14, 1949, remained high through the summer (average GSI from 6.6 to 12.5%), and decreased in late August back to very low levels. In contrast, black crappie increased their GSI during late summer and fall, with the level in November similar to the level the next March, hence, based on gonad size, he speculated that their gonads are ready to spawn with little additional energy needed. What determines the seasonal pattern of allocation to reproduction? Bunnell and Marschall (2003) combined dynamic programming with an IBM to examine optimal timing of energy allocation to reproduction in white crappie. The dynamic programming model for largemouth bass developed by Garvey and Marschall (2003) suggests that fish should begin storing energy for reproduction in the fall to prepare for reproduction in early spring. The costs of gonad development depend on the size of the gonads and their energy content. Female ovaries are much larger than male testes (James 1946). Ovary size varies with available energy. Breck’s (1996) pond experiments with bluegill showed that female GSI in June varied inversely with stocking density for fish stocked in April, but the number of eggs per gram of bluegill ovary was not significantly different among the stocking-density treatments. Energy content of ovaries is higher than the whole-body average, due to the relatively large amount of lipid in eggs. Bunnell and Marschall (2003) reported that ovaries of white crappie are about 1.33 times the energy density of somatic tissue. For bluegill and pumpkinseed, eggs in the ovary can be in different stages of development, and an individual female can deposit eggs at several times during the late spring and summer (James 1946; Fox and Crivelli 1998). This is in contrast to largemouth bass, in which eggs tend to develop at the same time and females deposit all their eggs in a short period (James 1946).
7.7.2 Nest building and guarding Nests are made by males using strong movements of the caudal fin directed downward toward the sediment, combined with compensatory movements by the pectoral fins to minimize forward motion of the fish. When a nest is built in very shallow water, some of the energy directed at excavating the nest is lost in making waves on the surface of the water. To our knowledge, the energy costs of nest building have not been measured. In all centrarchids, males guard nests from spawning through the swim-up stage. The period of guarding is of longer duration in black bass, because the males guard fry for several weeks after they have left the nest. Cooke et al . (2002) monitored male smallmouth and largemouth bass as they guarded nests at about 18◦ C and reported that the duration of stages at that water temperature was 3 days for the egg stage, 2 days for egg-sac fry, 2 days for swim-up fry, and 15 days for free-swimming fry, for a total of 22 days of guarding. For both smallmouth bass and bluegill, larger individuals tend to nest and spawn earlier than smaller adults and tend to be in better condition than smaller bluegill (Ridgway et al . 1991; Baylis et al . 1993; Lukas and Orth 1995; Cargnelli and Gross 1997; Wiegmann et al . 1997). A similar pattern has been observed for largemouth bass in pond experiments (Goodgame and Miranda 1993), but was not observed in Ohio reservoirs (Garvey et al . 1988). Energy costs of guarding are due to lack of feeding, and increased respiration rate due to increased activity. Most of the cost of nest guarding appears to be due to the foregoing of foraging. Male centrarchids do not leave the nest to forage in order to protect eggs and fry, which can be attacked by nest predators within seconds or minutes. This means that, apart
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from rare opportunistic feeding at the nest (Hinch and Collins 1991), the male must survive the guarding period using stored energy. Cooke et al . (2002) monitored the activity level of guarding black bass using radio transmitters with implanted electrodes that indicated activity of axial red muscles. They found that nesting males had elevated activity compared to nonnesting bass, that activity was elevated during both day and night, and that activity level varied with developmental stage of the brood. For example, for male largemouth bass, they determined average swimming activity of about 43 km d−1 (0.50 m s−1 ) during their offspring’s egg stage, 58 km d−1 (0.67 m s−1 ) during the egg-sac fry stage, 37 km d−1 (0.43 m s−1 ) during the swim-up fry stage, and 52 km d−1 (0.60 m s−1 ) during the free-swimming fry stage. Relatively long periods of low activity, including fanning the eggs, are interrupted by short periods of high activity that indicate rapid movements to drive away potential nest predators. They estimated that the activity level of nesting males was about double than that of nonnesting fish. The instantaneous activity level for free-swimming, nonnesting smallmouth bass was estimated to be 0.317 m s−1 , based on a similar study conducted in Lake Erie. Cooke et al . (2002) used the Wisconsin Bioenergetics Model 3.0 with parameters from Rice et al . (1983) to simulate energy expenditure of 1000 g nesting and nonnesting largemouth and smallmouth bass. They used activity levels that varied with the developmental stage of the offspring, as measured in the field. Over a 22-day nesting period, and accounting for variations in male activity level among brood developmental stages, they estimated that a nesting, nonfeeding largemouth bass would lose 125 g, whereas a nonnesting, nonfeeding male would lose 83 g in 22 days. A nesting, nonfeeding smallmouth bass would lose 115 g, whereas a nonnesting, nonfeeding male would lose 98 g. Based on these values, 66% of the mass lost by nesting largemouth bass is due to foregoing feeding, and 34% is due to increased activity. For nesting smallmouth bass, 85% of the mass lost is due to foregoing feeding, and 15% is due to increased activity. Costs of reproduction are high enough that males of most centrarchid species probably do not renest in the same year. In some systems, the cost of reproduction can significantly reduce survival of male smallmouth bass. Ridgway and Shuter (1994) provided supplemental food to parental smallmouth bass and found that this improved male survival in 1 year, compared to unfed males; in the second year it improved care duration and reproductive success but decreased survival. Wiegmann and Baylis (1995) had evidence that few male smallmouth bass were repeat spawners in the northern lake they studied. Apparently, the cost of reproducing was sufficiently high that the population they were studying was nearly semelparous, with up to 94% of males dying after their first reproduction.
7.7.3 Egg development time Egg development time can be included in an energetics model of a centrarchid full life cycle. Egg development time is also important in determining the duration of male parental care, thereby affecting the cost of parental care and the duration of high vulnerability to angling. Trebitz (1991) developed the following equation for time to hatch (th , day) for eggs of largemouth bass as a function of average water temperature (Tave ,◦ C), based on data in Heidinger (1976). e9.88 T −1.88 24 Gillooly et al . (2002) recently derived a model from first principles of allometry and biochemical kinetics that explains the general pattern across taxonomic groups: egg development time is predominantly a function of temperature and egg size. The time to hatch is shorter at higher temperatures and for smaller eggs. Their model explained 83% of the variance in a data set of 59 values for fish egg size, temperature, and time to hatch (Gillooly and Dodson 2000; Gillooly et al . 2002, their Figure 1). Their general model was fitted to data for both aquatic ectotherms (including fishes and amphibians) and birds: 4m1/4 th = a(Tc ) where th is the time (day) to hatch, m is the egg mass (g), and a(Tc ) is a function that expresses the temperature dependence of development: αTc + yint a(Tc ) = 4 exp − 1 + Tc /T0 th =
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40 2.00 mm, smallmouth bass 1.67 mm, largemouth bass Time to hatch (d)
30
1.45 mm, redear sunfish 1.20 mm, bluegill 1.05 mm, pumpkinseed
20
0.89 mm, white crappie 10
0 10
15
20
25
30
Temperature (°C) Figure 7.8 Predicted time to hatch (days) increases with egg diameter (millimeter, converted to mass, gram) and decreases with temperature (Gillooly et al . 2002). Egg diameters are representative of several common centrarchid species.
where α = −0.12 per degree Celsius is the slope, and yint = 6 is the intercept of the regression of mass-adjusted time to hatch versus scaled temperature, T0 = 273 Kelvin is the freezing point of water, and Tc is the temperature (◦ C) to which the eggs are exposed. Gillooly et al . (2002) also indicate how to compute the time to hatch for fluctuating temperatures. Winemiller and Rose (1992) compiled life history characteristics for many North American fishes, including 16 species of centrarchids. In their compilation, mean ovum size in centrarchids varies from 0.89 mm in white crappie, to 1.20 mm in bluegill, to 2.00 mm in smallmouth bass. According to the model of Gillooly et al . (2002), the egg-size effect would result in faster development times in white crappie than smallmouth bass, with bluegill intermediate. Because smallmouth bass spawn at cooler temperatures (15 or 16◦ C) than bluegill (about 20◦ C), the temperature effect would result in faster development times for bluegill. Figure 7.8 shows the predicted effect of temperature and egg size on time to hatch for several egg sizes, selected to represent the range of centrarchid egg sizes in the compilation by Winemiller and Rose (1992). These predicted hatching times, based on the broad biological pattern across many taxa, are probably overestimates for centrarchids. However, the general relationships presented may encourage others to measure these data and estimate α and yint specifically for centrarchids and consider the possible consequences of egg mass, temperature, and hatching time for evolution of life history traits, including the trade-off between egg size and egg number (Gillooly and Dodson 2000).
7.7.4 Modeling energetics of reproduction Modeling energetics of reproduction has grown increasingly sophisticated, particularly in addressing questions involving reproduction. In early bioenergetics models, including the first and second versions of the Wisconsin software package (Hewett and Johnson 1987, 1992), fish biomass (W ) was the only state variable, and reproduction was incorporated by decrementing W once per year by the average reproductive loss (G). For example, on the specified annual date of reproduction, if average reproductive losses were 10% of body mass, then W would be decremented by G = 0.10 ∗ W . Using this same basic approach, separate model runs can be done for males and females, to account for differences in gonad mass lost at spawning. Adjustments can also be made to account for the energy density of eggs and milt relative to that of the rest of the body. The standard model can be used to estimate the costs of male guarding of offspring, by increasing fish metabolic rate (by increasing the simulated swim speed) and setting consumption to zero during the guarding period. Cooke et al . (2002) used this approach to estimate the energy costs of parental care by male largemouth bass and smallmouth bass, as described earlier.
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Bunnell and Marschall (2003) used a dynamic programming approach to determine optimal allocation of energy between growth in length and growth of gonads during a year for different feeding conditions. Length and gonad mass were tracked separately for the fish, a change from the Wisconsin model. The optimal allocation pattern was the one that maximized lifetime fitness of modeled fish, with fitness evaluated as the expected number of larvae produced. Their analysis of white crappie in Ohio reservoirs indicated that some energy should be allocated to gonad development as early as fall if feeding conditions might not be favorable in the spring. Interestingly, the benefit of allocating energy to ovary development in the fall was strongly dependent on the likely feeding conditions the following spring, not on feeding conditions earlier in the summer. Garvey and Marschall (2003) developed a model to analyze energy allocation patterns of largemouth bass for different feeding conditions and different latitudes. Their model separately tracked allocation of energy to somatic growth (increase in length), fat, and reproductive tissue. Their question required the model to include these separate compartments. A theoretical reciprocal transplant experiment allowed them to compare the success of fish with an optimal northern strategy “transplanted” to a southern environment versus the success of fish with an optimal southern strategy “transplanted” to a northern environment. The southern strategy was not successful in the north because they did not allocate sufficient energy to fat reserves prior to winter and did not begin early enough to allocate energy to ovary development.
7.8 Synthesis Because rapid growth and high reproductive output are highly selected for in most fishes, behaviors that result in maximizing net energy gain should be commonly observed. Such behaviors include both habitat selection and feeding choices. One feature of habitat choice that has been described for several centrarchid species is temperature preference (see related discussion in Chapter 8 of this book). Fish typically prefer a narrow range of temperatures within which physiological processes, such as digestion, energy assimilation, and growth, are presumed to be most efficient (Reynolds 1977; Magnuson and Beitinger 1978; Jobling 1981). Several laboratory studies have demonstrated the ability of centrarchid species (bluegill, rock bass, black crappie, smallmouth bass, and largemouth bass) to maintain their temperatures within a few degree range in shuttle-box test chambers (Neill and Magnuson 1974; Reynolds and Casterlin 1978; Wildhaber and Crowder 1990). Centrarchid species have also been observed exhibiting thermoregulatory behavior in the field (Zimmerman et al . 1989). As with choice of thermal habitat, the type of feeding strategy exhibited also has energetic consequences. Several investigators have used centrarchid species as experimental subjects to test predictions of optimal foraging theory. Werner (1974) and Werner and Hall (1974) used theoretical modeling and laboratory experimentation with bluegill to show that prey size selection is related to the optimal allocation of time spent searching for and handling prey. Stein (1977) similarly found that smallmouth bass size-selectivity on crayfish prey resulted in maximizing net energy intake by balancing handling and pursuit time with prey caloric content. To the contrary, Stein et al . (1984) found that prey selection by redear sunfish on various snail species in the laboratory was not consistent with optimal foraging predictions. Crowder and Cooper (1982) and Savino et al . (1992) both studied the relationship between vegetation density and the ability of bluegill to choose prey that maximized net energy intake. The experimental results of Savino et al . (1992) did not find a difference in growth rate (i.e. net energy intake) at different plant densities as predicted by Crowder and Cooper (1982). Manatunge and Asaeda (1999) developed a model to predict the feeding selectivity of white crappie and bluegill based on the caloric benefit and energetic cost of searching for and consuming different sized zooplankton. A comparison of stomach samples from fish in the field to the available prey base at the same location demonstrated that the fish were selecting prey as predicted by the model, which matched predictions of optimal foraging theory. Living an energetically efficient life in the wild, however, is rarely as simple as selecting the right temperature or consuming the perfect size prey. More often than not these choices are entangled in consideration of several factors in combination. For example, Neil and Magnuson (1974) tested the effect of food availability on behavioral thermoregulation and found that bluegill would spend increased time at temperatures above the preferred temperature to acquire food. Bevelhimer (1996) found similar results in laboratory studies where smallmouth bass altered the amount of time they spent at higher than preferred temperatures depending on food availability. Smallmouth bass were willing to spend significant time at an energetically costly temperature in exchange for foraging opportunities, especially when the available ration was less than satiation; when satiated, smallmouth bass returned to cooler, lower cost temperatures.
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Energetic trade-offs also occur in habitat choices related to things other than food availability. Energy efficiency is sometimes compromised for survival as shown in studies that demonstrated how risk of predation can cause fish to exhibit less than optimal feeding strategies (Werner et al . 1983; Mittelbach 1984). Similarly, Bevelhimer (1995, 1996) demonstrated that, in both field and laboratory settings, smallmouth bass would forego habitats with more energetically favorable thermal conditions in favor of those with physical cover and suboptimal temperatures. Because the standard bioenergetics models are not able to handle the energetic consequences of predator avoidance and many habitat choices, some researchers have instead used IBMs mentioned earlier to incorporate factors that directly and indirectly affect fish energetics. These models have been used to investigate smallmouth bass YOY recruitment (DeAngelis et al . 1991), largemouth bass juvenile survival and recruitment (Trebitz 1991), bluegill fry density and growth relationships (Breck 1993), and the effects of alternate river flow regimes on smallmouth bass populations (Jager et al . 1993).
7.9 Research needs Like many other fundamental aspects of fish biology, basic studies on fish energetics seem to have become pass´e to many researchers. Analytical methods and instruments that can be used to measure various aspects of fish physiology and biochemistry are no doubt much more advanced than 10 or 20 years ago, but the volume of empirical studies on fish energetics seems to not have kept up with the technology. Basic energetic relationships have not been determined for most centrarchid species. In the absence of species-specific data for parameters used in bioenergetics models, researchers resort to borrowing parameters from other species to construct new models. For closely related species with similar life histories (e.g., diet, body shape, and behavior), one might expect to find similar physiological energetics. Unfortunately, few simultaneous studies have been performed with multiple species to determine the degree of similarity among congeneric species. The only such study we found for centrarchids is O’Hara (1968) who determined metabolic rates for bluegill and pumpkinseed sunfish. A good example of such a study is Bevelhimer et al . (1985) who simultaneously determined rates of metabolism, food consumption, and growth for two esocid species and a hybrid. Along these lines, there is a great need for detailed studies on the bioenergetic differences between largemouth bass and Florida bass given the economic and recreational importance of these species. Other than for largemouth bass, there are no studies on geographic variation in bioenergetics parameters within other species. Virtually nothing is known of the energetics of the smallest centrarchids, such as the Enneacanthus species. There are several aspects of fish energetics that are still poorly understood, not just for Centrarchidae but for other families as well. General areas in need of more research include costs associated with activity and reproduction, allometry of feeding and respiration (i.e. the size-dependent coefficients in explanatory equations), and the effects of ration or nutritional status on energetic processes including compensatory growth. For example, the need for improvement in the ability of a largemouth bass model to handle fish of different sizes was demonstrated in a study with juvenile largemouth bass by Slaughter et al . (2004). Recent advances in understanding compensatory growth of centrarchid species are promising and have significant relevance to fish biology in general as discussed earlier. Of the 33 centrarchid species, we found published bioenergetics models for only 5 species (see Table 7.1; we did not count the spotted bass model which was a copy of a smallmouth model or the hybrid sunfish model which was a copy of a bluegill model). This area of research is wide-open for further development. Unfortunately, the development of models for new species usually depends on some management or conservation need and not so much on a basic quest to fill gaps in knowledge. An alternative to developing individual models for a variety of similar species would be the development of a reliable generic model that could be used interchangeably. Simultaneous laboratory studies would first be needed to define the degree of similarity among different genera and species. For some model parameters, differences among species are either nonexistent or so small that the same value may be representative of all species. For parameters where significant differences are observed, it might be possible to use adjustable parameters that are applied automatically based on various life history parameters, such as shape, diet, feeding mode, maximum size, level of activity, latitudinal distribution, etc. The general shape and magnitude of the relationships between some parameters (e.g., metabolic rate and feeding rate) and temperature may be applicable to several species by simple adjustment to the left or right along the temperature axis to account for
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different thermal optima. A comprehensive comparison of bioenergetics parameters for centrarchid species to those of other families (such as salmonids) would provide useful insight into the degree of within-family differences relative to amongfamily differences. Such an analysis would certainly add perspective to the importance or lack of species-specific models. More model validation and testing are desperately needed. A significant deficiency in bioenergetics modeling to date for all species has been the scarcity of studies with independent evaluations. Depending on the type of application, the level (or lack thereof) of model validation dictates the level of confidence one can have in the results. The greater the number of parameters borrowed from other studies, stocks, or species, the greater the need for model validation and testing. This is also the case when models are applied beyond the conditions (e.g., temperature, fish size, and ration), under which parameters were derived. Due to the lack of that one comprehensive and fully tested study that defines all the energetic parameters for a single species, there are many scientists and managers that will continue to view bioenergetics models with a great deal of suspicion. Centrarchid species, perhaps second only to salmonids in North America, have been the subject of a plethora of physiological studies and bioenergetics analyses for many years. The knowledge base derived from these studies has made possible the development of energy budgets and bioenergetics models that have been used to address a variety of fisheries management and conservation questions. The bioenergetics approach to investigating fish behavior, foraging, and growth is based on solid theory and has been proven to be a sound approach in fisheries science. However, our knowledge of fish energetics and our application of that knowledge in the form of bioenergetics models are still fraught with uncertainty and probably at least a little misunderstanding. As we continue to use bioenergetics models, we must also continue performing the basic experiments to continue to refine the underlying knowledge base. When specific metabolic functions and relationships become better understood and bioenergetics models are continually tested and improved, we can expect the use of such models and other energetic-based approaches to become more common and more useful in the management and conservation of centrarchid species.
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Whitledge, G. W., P. G. Bajer, and R. S. Hayward. 2006b. Improvement of bioenergetics model predictions for fish undergoing compensatory growth. Transactions of the American Fisheries Society 135: 49–54. Wiebe, A. A. and A. C. Fuller. 1933. The oxygen consumption of largemouth black bass (Huro floridana) fingerlings. Transactions of the American Fisheries Society 63: 208–214. Wiegmann, D. D. and J. R. Baylis. 1995. Male body size and paternal behavior in smallmouth bass Micropterus dolomieui (Pisces: Centrarchidae). Animal Behaviour 50: 1543–1555. Wiegmann, D. D., J. R. Baylis, and M. H. Hoff. 1997. Male fitness, body size and timing of reproduction in smallmouth bass, Micropterus dolomieui . Ecology 78: 111–128. Wildhaber, M. L. and L. B. Crowder. 1990. Testing a bioenergetics-based habitat choice model: bluegill (Lepomis macrochirus) responses to food availability and temperature. Canadian Journal of Fisheries and Aquatic Sciences 47: 1664–1671. Wildhaber, M. L. and P. J. Lamberson. 2004. Importance of the habitat choice behavior assumed when modeling the effects of food and temperature on fish populations. Ecological Modelling 175: 395–409. Winberg, G. G. 1956. Rate of metabolism and food requirements of fishes. Fisheries Research Board of Canada Translation Series No. 194, 1960, Ottawa, Canada. Windell, J. T. 1966. Rate of digestion in the bluegill sunfish. Investigations of Indiana Lakes and Streams 7: 185–214. Winemiller, K. O. and K. A. Rose. 1991. Patterns of life-history diversification in North American fishes: implications for population regulation. Canadian Journal of Fisheries and Aquatic Sciences 49: 2196–2218. Wohlschlag, D. E. and R. O. Juliano. 1959. Seasonal changes in bluegill metabolism. Limnology and Oceanography 4: 195–209. Wright, R. A., J. E. Garvey, A. H. Fullerton, and R. A. Stein. 1999. Predicting how winter affects energetics of age-0 largemouth bass: how do current models fare? Transactions of the American Fisheries Society 128: 603–612. Wright, R. M. 1990. Aspects of the ecology of bream, Abramis brama (L.), in a gravel pit lake and the effects of reducing the population density. Journal of Fish Biology 37: 629–634. Xie, S., X. Zhu, Y. Cui, R. J. Wootton, W. Lei, and Y. Yang. 2001. Compensatory growth in the gibel carp following feed deprivation: temporal patterns in growth, nutrient deposition, feed intake and body composition. Journal of Fish Biology 59: 999–1009. Yako, L. A., M. E. Mather, and F. Juanes. 2000. Assessing the contribution of anadromous herring to largemouth bass growth. Transactions of the American Fisheries Society 129: 77–88. Zimmerman, L. C., E. A. Standora, and J. R. Spotila. 1989. Behavioural thermoregulation of largemouth bass (Micropterus salmoides): response of naive fish to the thermal gradient in a nuclear reactor cooling reservoir. Journal of Thermal Biology 14: 123–132. Zweifel, R. D. 2000. Development and evaluation of a bioenergetics model for white crappie. Master’s thesis. University of Missouri, Columbia, MO. Zweifel, R. D., R. S. Hayward, and C. F. Rabeni. 1999. Bioenergetics insight into black bass distribution shifts in Ozark border region streams. North American Journal of Fisheries Management 19: 192–197.
Chapter 8
Physiology and organismal performance of centrarchids J. D. Kieffer and S. J. Cooke
8.1 Introduction The relationship between fish and their environment represents some of the most fascinating examples of physiological adaptations among the animal kingdom. This is particularly the case for temperate fishes that experience distinct seasons that can include both harsh and cold winters and hot summers. With a few exceptions, most fish are ectotherms and therefore, their physiological processes and organismal performance, including swimming ability, metabolic rates, and enzyme activity are often dictated by water temperature. Furthermore, many freshwater fish live in a heterogeneous environment where oxygen concentrations vary extensively, making aerobic respiration a challenge. Individuals, populations, and species exhibit different tolerances to environmental conditions such as hypoxia (low oxygen) and temperature. When exposed to stressors, be it anthropogenic (e.g., angling, handling) or natural (e.g., winter, predation attempt, hypoxia), fish also exhibit variation in how they respond. Much of the existing research on the physiology, performance, and environmental tolerances of fishes involves the use of a salmonid model, and most frequently the rainbow trout. Aside from several other unique examples, such as tuna, Arctic fish, and desert fish, that have been well studied because they live in extreme environments or have special adaptations, we know less about the physiology of other fish groups, such as the centrarchids (sunfish). However, centrarchids have been studied because of their use in culture facilities, their hardiness in the laboratory, their wide availability in natural systems, and their importance in recreational fisheries. In addition, centrarchid fishes have value for use as physiological models because of their large variation in life history, both within and among species. Centrarchid fishes occupy a number of different niches and habitats and vary extensively in body size. The behavioral ecology and natural history of several species of centrarchids (e.g., bluegill, Lepomis macrochirus, largemouth bass, Micropterus salmoides) have been well studied (Miller 1975; Keast 1978; Gross 1980) such that this information can be used to form interesting hypotheses. Despite this background information and apparent value in studying centrarchid biology, there is a relative paucity of research on the physiology and performance of these fishes. In fact, most accounts on this group of fishes are rather disparate with no cohesive synthesis on an individual species or the family. For this reason, it has not been possible to develop a more general understanding of centrarchid physiology and performance. For the first time, we synthesize existing information on the physiology of centrarchid fishes. The specific objective of this chapter is to provide a detailed overview of the physiology of centrarchid fishes, with particular emphasis on how they cope with environmental (e.g., temperature, hypoxia) and biological (e.g., swimming, exercise) challenges. We adopt a comparative approach and discuss the relevance of centrarchid physiology to ecology, behavior and, when appropriate, management. Because of the wealth of knowledge on the physiology of salmonids, we also provide some direct comparisons between the physiology and performance of those fishes and the centrarchids. We conclude by providing a general discussion of the physiological patterns evident among the centrarchids focusing on both general patterns and variation. Issues that are applied are also covered in detail in other chapters (e.g., swimming performance relative to fishway design; sublethal impacts of catch-and-release angling). Other information relative to fish physiology and performance can also be found in the chapter of this volume on bioenergetics (Chapter 7).
207
208
Centrarchid fishes
8.2 Baseline physiological variables A compilation of baseline physiological values for oxygen consumption rates, ventilation rates, blood variables, such as ion concentrations, energy reserves (e.g., protein and glucose), plasma hormone concentrations, and muscle enzyme and metabolite levels of various centrarchids is found in Table 8.1. We have also included in the table information about the size of the fish and the water temperature. As with any data compilation, it is important to recognize that factors such as sampling method and sampling time can significantly affect baseline physiology. McDonald and Milligan (1992) discuss some of these general issues and provide an excellent review of the chemical properties of the blood of fish (mainly salmonids). From Table 8.1, it is interesting to note that most of the baseline physiological data exist for only a few of the 32 species of centrarchid fishes. In fact, for more than 20 of the centrarchid species, there is simply no published physiological data at all. As noted for other species of fish [e.g., rainbow trout, Kieffer et al . (1998); tilapia, Oreochromis niliticus, Alsop et al . (1999); sockeye salmon, Oncorhynchus nerka and coho salmon, Oncorhynchus kisutch, Lee et al . (2003a)], temperature strongly affects baseline oxygen consumption rates in various centrarchids (see Figure 8.1a). In addition to these temperature related effects, there are differences in oxygen consumption rates between species of centrarchids (Figure 8.1a). From the available data, bluegill (L. macrochirus) and longear sunfish (Lepomis megalotis) appear to have higher resting metabolic rates across temperatures relative to largemouth bass and pumpkinseed sunfish (Lepomis gibbosus). Whether this variation represents true species differences or is the result of different methodologies is not certain. However, there are clear differences in routine oxygen consumption rates across temperatures among different stocks of largemouth bass (Cooke et al . 2001a; see Figure 8.1b). These intraspecific findings have also been noted for Arctic charr (Salvelinus alpinus) (Giles 1991). Overall, the values noted for centrarchids are in line with those of other species, such as sockeye and coho salmon (Brett 1995; Lee et al . 2003a) and brown trout (Salmo trutta) (Sloman et al . 2000). In addition to temperature, baseline oxygen consumption rates are also significantly influenced by the levels of oxygen in the water (Cech et al . 1979; Figure 8.1c). These effects are particularly noticeable at higher (i.e. 30◦ C) temperatures (Figure 8.1c). Cech et al . (1979) discuss these findings with respect to oxygen levels required for largemouth bass (see Section 8.4.4 on hypoxia for more detail). Ventilation rates are variable among centrarchid species (26–75 beats min−1 ; Table 8.1). It appears that this variation is largely related to body size and water temperature. However, these rates are consistent with ventilation rates noted for other species of fish, including shortnose (Acipenser brevirostrum) and Atlantic (Acipenser oxyrhinchus oxyrhinchus) sturgeon (Baker et al . 2005) and rainbow trout (Oncorhynchus mykiss; Bindon et al . 1994). Blood hemoglobin and hematocrit (Hct) are within the normal ranges for other fishes (Table 8.1; reviewed in Gallaugher and Farrell 1998), suggesting that centrarchids have adequate oxygen transport capacities. Resting ion levels (Na+, K+, Cl− ) and osmolality (a measure of the total concentration of ions in the blood) are within the ranges noted for other freshwater fishes (McDonald and Milligan 1992) and are probably sensitive to a wide variety of abiotic (e.g., salinity, pollutants) and biotic (e.g., stress) influences (see Section 8.4). Of all the muscle and blood variables noted in Table 8.1, the greatest difference between centrarchids and other species is the muscle lactate dehydrogenase (LDH) concentration. This enzyme is important in the conversion of pyruvate to lactate during anaerobic metabolism. Overall, centrarchids have relatively low levels of LDH in their muscle relative to Atlantic salmon (Salmo salar) (McDonald et al .1998) and sea lamprey (Petromyzon marinus) (Boutilier et al . 1993, Wilkie et al . 2001). This lower LDH level (∼ one-fifth to one-tenth of the levels of Atlantic salmon and sea lamprey, depending on the temperature) would imply that the anaerobic capacity of centrarchids is significantly lower compared to other species. Indeed, this is the case as centrarchids generally produce less lactate than other species of fish following certain stress (e.g., exhaustive exercise, hypoxia; see Table 8.2).
8.3 Physiological challenges/tolerances 8.3.1 Swimming capacity For various practical and physiological reasons, most of the research on the swimming capacity in fish has focused on salmonids (Brett 1964, Davison 1989). Levels of swimming performance in fish are traditionally defined in terms of duration of swimming (Beamish 1978) and whether the exercise depends on the presence of oxygen (i.e. aerobic
Physiology and organismal performance of centrarchids
209
Table 8.1 Baseline physiological variables of various centrarchid fish species. Physiological variable and species Oxygen consumption Pumpkinseed
Pumpkinseed
Bluegill
Smallmouth bass
Variable specific units −1
mg kg
hr
Banded sunfish Longear sunfish
White crappie
Source
−1
8
100 g
Evans 1984
45
16
100 g
Evans 1984
45
20
45 g
Brett and Sutherland 1965
70
24
100 g
Evans 1984
81
28
100 g
Evans 1984
117
32
100 g
Evans 1984
∼40
15
MA Population
∼43
15
NC Population
Roberts 1967
∼70
25
MA Population
Roberts 1967
Roberts 1967
∼90
25
NC Population
Roberts 1967
111.9
10
9.74 cm (31 g)
Dent and Lutterschmidt 2003
143.6
20
10.3 cm (36 g)
100
25
118 g
223.4
30
9.83 cm (31 g)
86.7
–
32 cm
74.7 nmol g
Green sunfish
Size of fish (various units)
30
−1
Largemouth bass
Water temperature (◦ C)
min
−1
26.6
Dent and Lutterschmidt 2003 Marvin and Heath 1968 Dent and Lutterschmidt 2003 S. Peake, unpublished data
18
15 g
5
6–10 g/8.3–9.7 cm
Johnson and Charlton 1960
Gonzalez and McDonald 1994
91
12
6–10 g/8.3–9.7 cm
Johnson and Charlton 1960
93.8
17
6–10 g/8.3–9.7 cm
Johnson and Charlton 1960
129.5
22
6–10 g/8.3–9.7 cm
Johnson and Charlton 1960
224
29
6–10 g/8.3–9.7 cm
Johnson and Charlton 1960
∼78
10
150 g/22.5 cm
Beamish 1970
∼95
15
150 g/22.5 cm
Beamish 1970
∼110
20
150 g/22.5 cm
Beamish 1970
∼125
25
150 g/22.5 cm
Beamish 1970
∼150
30
150 g/22.5 cm
Beamish 1970
∼160
34
150 g/22.5 cm
67.7
20
230–470 g
Cech et al. 1979
102.9
25
230–470 g
Cech et al. 1979
173.2
30
230–470 g
Cech et al. 1979
130
15
4.9–15.8 g
41.3 nmol g−1 min−1
18
–
Beamish 1970
Horkel and Pearson 1976 Gonzalez and McDonald 1994
126.7
10
8.9 cm (22 g)
Dent and Lutterschmidt 2003
169.2
20
9.44 cm (30 g)
Dent and Lutterschmidt 2003
187.7
30
9.36 cm (25 g)
Dent and Lutterschmidt 2003
121.5
25
∼90 g
Parsons and Sylvester 1992 (continued)
210
Centrarchid fishes
Table 8.1 (continued). Physiological variable and species Ventilation rate Green sunfish Largemouth bass Smallmouth bass Bluegill Pumpkinseed
Variable specific units vents min
−1
Water temperature (◦ C)
Size of fish (various units)
Source
–
–
26
15
6–10.5 cm/5–16 g
∼65
20
422 g
∼120
20–22
16–20 cm
∼75
20
389 g
Furimsky et al. 2003
70
25
118 g
Marvin and Heath 1968
∼40
15
MA Population
Horkel and Pearson 1976 Furimsky et al. 2003 C. Suski, unpublished data
Roberts 1967
∼25
15
NC Population
Roberts 1967
∼50
25
MA Population
Roberts 1967 Roberts 1967
∼52
25
NC Population
G dl−1
–
–
Bluegill
9.07
20–22
∼33 g
Heath 1991
Largemouth bass
8.1
–
115 g
Black 1955
6.2
–
1–4 yr
5.72–7.87
over seasons
–
White crappie
6.1
22
7.5 cm
Rock bass
5.85
–
–
7
9
14.2–18.4 cm
Bidwell and Heath 1993 Bidwell and Heath 1993
Hemoglobin
Redear sunfish
6.5 Hematocrit (%) Bluegill
Smallmouth bass
Largemouth bass
7
14.2–18.4 cm
–
–
Clark et al. 1979 Atkinson and Judd 1978 Parsons 1993 Coburn 1970 in Atkinson and Judd 1978
30%
23
–
Musselman et al. 1995
35%
–
–
Coburn 1970 in Atkinson and Judd 1978
37%
20
5–7 cm
Lemly 1993
25%
–
32 cm
S. Peake, unpublished data
19.20%
20
–
Furimsky et al. 2003
33%
12, 16, and 20
–
J. Schreer, unpublished data
35%
–
1–4 yr
16.70%
20
–
Furimsky et al. 2003
∼40%
over seasons
–
Atkinson and Judd 1978
31%
–
–
Coburn 1970 in Atkinson and Judd 1978
27%
22
7.5 cm
Black crappie
23%
–
–
Coburn 1970 in Atkinson and Judd 1978
Rock bass
30%
–
–
Coburn 1970 in Atkinson and Judd 1978
Redear sunfish White crappie
Clark et al. 1979
Parsons 1993
35%
9
14.2–18.4 cm
Bidwell and Heath 1993
32%
7
14.2–18.4 cm
Bidwell and Heath 1993 (continued)
Physiology and organismal performance of centrarchids
211
Table 8.1 (continued). Physiological variable and species Plasma sodium
Variable specific units
Size of fish (various units)
Source
–
–
Bluegill
∼144
20–22
∼33 g
Heath 1991
Smallmouth bass
∼173
22
25 cm
Carmichael et al. 1983
149
12, 16, and 20
–
J. Schreer, unpublished data
Largemouth bass
125
–
–
Suski et al. 2003
Plasma chloride
mequiv l−1
–
–
∼110
20–22
∼33 g
112
22
25 cm
101
12, 16, and 20
–
Bluegill Smallmouth bass Largemouth bass
mequiv l
−1
Water temperature (◦ C)
∼100
–
–
101–115
–
150 g
Heath 1991 Carmichael et al. 1983 J. Schreer, unpublished data Suski et al. 2003 Williamson and Carmichael 1986
106
23
13–23 cm
mequiv l−1
–
–
Bluegill
∼5.5
20–22
∼33 g
Smallmouth bass
∼2.5
22
25 cm
2.99
12, 16, and 20
–
J. Schreer, unpublished data
∼2.5
–
–
Suski et al. 2003
MOsm
–
–
Plasma potassium
Largemouth bass Plasma osmolarity Rockbass
Carmichael et al. 1984a Heath 1991 Carmichael et al. 1983
325
9
14.2–18.4 cm
Bidwell and Heath 1993
∼280
7
14.2–18.4 cm
Bidwell and Heath 1993
293
–
3.2 g
McCormick et al. 1989
Bluegill
∼300
23
–
Musselman et al. 1995
Largemouth bass
∼300
22–24
33 cm
286
23
5–6 cm
294
23
13–23 cm
g dl−1
–
–
Black crappie
Plasma protein Rock bass
Suski et al. 2004 Susanto and Peterson 1996 Carmichael et al. 1984a
6
9
14.2–18.4 cm
∼5.5
7
14.2–18.4 cm
Largemouth bass
7
–
1–4 yr
Clark et al. 1979
6.97
–
51–2382 g
Clark et al. 1979
Plasma glucose
mg dl−1
–
–
Largemouth bass
Smallmouth bass
100
–
–
161.2
–
1–4 yr
53
23
13–23 cm
68–74
–
150 g
157.8
N/A
51–2382 g
∼90
22
25 cm
48
12, 16, and 20
–
Bidwell and Heath 1993 Bidwell and Heath 1993
´ Sepulveda et al. 2004 Clark et al. 1979 Carmichael et al. 1984a Williamson and Carmichael 1986 Clark et al. 1979 Carmichael et al. 1983 J. Schreer, unpublished data (continued)
212
Centrarchid fishes
Table 8.1 (continued). Physiological variable and species
Variable specific units
Water temperature (◦ C)
Size of fish (various units)
Bluegill sunfish
∼50
20–22
∼33 g
Rockbass
∼50
9
14.2–18.4 cm
Bidwell and Heath 1993
∼50
7
14.2–18.4 cm
Bidwell and Heath 1993
Source Heath 1991
Muscle cytochrome oxidase
umol min−1 g−1
White crappie
∼5
5
–
Tschantz et al. 2002
∼8
25
–
Tschantz et al. 2002
Black crappie
∼35
25
–
Tschantz et al. 2002
Bluegill sunfish
∼3
5
–
Tschantz et al. 2002
∼1
25
–
Tschantz et al. 2002
∼2
5
–
Tschantz et al. 2002
∼4
25
–
Tschantz et al. 2002
∼2
5
–
Tschantz et al. 2002
∼2
25
–
Tschantz et al. 2002
Green sunfish Largemouth bass Muscle lactate dehydrogenase White crappie Black crappie Bluegill sunfish Green sunfish Largemouth bass
umol min−1 g−1 ∼30
5
–
Tschantz et al. 2002
∼75
25
–
Tschantz et al. 2002
∼25
5
–
Tschantz et al. 2002
∼100
25
–
Tschantz et al. 2002
∼25
5
–
Tschantz et al. 2002
∼40
25
–
Tschantz et al. 2002
∼38
5
–
Tschantz et al. 2002
∼80
25
–
Tschantz et al. 2002
∼38
5
–
Tschantz et al. 2002
∼100
25
–
Tschantz et al. 2002
Muscle phosphocreatine PCr
umol g−1
Largemouth bass
∼3
–
–
∼15
22–24
33 cm
Suski et al. 2005
Smallmouth bass Muscle ATP Largemouth bass
Smallmouth bass
Dehn and Schirf 1986
∼16
–
–
Suski et al. 2003
∼15
16
32 cm
Kieffer et al. 1995
−1
umol g ∼4
–
–
∼7
22–24
33 cm
Suski et al. 2005
∼7
–
–
Suski et al. 2003
∼6
16
32 cm
Kieffer et al. 1995
Dehn and Schirf 1986
(continued)
Physiology and organismal performance of centrarchids
213
Table 8.1 (continued). Physiological variable and species Plasma cortisol Largemouth bass
Plasma adrenaline
Variable specific units ng ml
Water temperature (◦ C)
Size of fish (various units)
Source
−1
16
21
–
Davis and Parker 1986
∼20
–
–
Suski et al. 2003
∼50
22–24
33 cm
Suski et al. 2005
nmol l−1
Largemouth bass
1
20
–
Furimsky et al. 2003
Smallmouth bass
∼1
20
–
Furimsky et al. 2003
Plasma noradrenaline
nmol l
−1
Largemouth bass
∼6
20
–
Furimsky et al. 2003
Smallmouth bass
∼2
20
–
Furimsky et al. 2003
Note: Values may be estimated.
versus anaerobic). Researchers have categorized swimming in fish into three broad categories: sustained, prolonged, and burst-type swimming (see Brett 1964; Beamish 1978; Hammer 1995; Plaut 2001, for details). Sustained exercise is powered exclusively by aerobic metabolism and, in fish, this type of exercise is referred to as cruising. Sustained swimming performance includes those speeds that can be maintained for long periods of time (typically greater than 200 min) without resulting in muscular fatigue (Beamish 1978). Prolonged exercise can last between 2 and 200 minutes and, depending on the swimming speed, is terminated by exhaustion. A third type of exercise, intense burst activity, relies almost exclusively on anaerobic metabolism within the white muscle and can only be maintained for short periods of time (typically less than 20 s). This type of exercise results in a significant reduction of intracellular energy supplies or by the accumulation of waste products (Kieffer 2000).
8.3.2 Tests to measure swimming performance in fish (methodological approaches) Different methods have been developed to quantify exercise performance in fish. Common laboratory tests include: (i) fixed velocity (fatigue) tests and (ii) increased (incremental) velocity tests (Brett 1964; Beamish 1978; Hammer 1995, for a review). Fixed velocity tests involve placing fish in a swim tunnel (or long raceway) and after an adjustment period, swimming speed is increased (steadily, in small steps, or abruptly) until the test velocity is achieved, after which the velocity is constant. At this point, time to swimming failure at the test speed is measured. An increased velocity test (also known as the UCrit, or the critical velocity test; Brett 1964) involves forcing fish against a known current. Unlike that of the fixed velocity tests, fish are exposed to increasing velocity increments (e.g., 10 cm s−1 ) and duration (ranges from 5 to 60 min; see Farlinger and Beamish 1977) until exhaustion is reached (see Brett 1964; Farlinger and Beamish 1977; Beamish 1978; Hammer 1995; Kolok 1999, for details and methodological considerations). Much of the earlier work on swimming in centrarchids has been documented in an excellent review by Beamish (1978). Recent research on swimming performance in centrarchids has focused on critical swimming capacity and fixed velocity performance. Some research has also focused on the metabolic costs (i.e. oxygen consumption) associated with swimming (see later).
8.3.3 Critical swimming (UCrit) performance Critical swimming speeds [UCrit, typically measured as body lengths (BLs) per second, BL s−1 ] for many species of centrarchid fish, such as largemouth bass, smallmouth bass (Micropterus dolomieu), pumpkinseed sunfish, bluegill sunfish
214
Centrarchid fishes
(a)
250 200 150 100 50 0 0
5
10
15
20
25
30
35
Oxygen consumption (mg O2 Kg−1 h−1)
Temperature (°C) (b)
200 160 120 80 40 0
6
12
18
Temperature (°C) (c)
160
30°C
120 25°C 80 20°C 40 0 30
40
50 60 70 80 90 100 110 120 130 140 150 160 Partial pressure of oxygen in water (mmHg)
Figure 8.1 (a) Baseline metabolic rates of various centrarchids as a function of water temperature. Closed circles represent longear sunfish (Dent and Lutterschmidt 2003), closed squares represent bluegill sunfish (Marvin and Heath 1968; Dent and Lutterschmidt 2003), open triangles represent largemouth bass (Johnson and Charlton 1960), open circles represent largemouth bass (Beamish 1964) and closed diamonds represent pumpkinseed sunfish (Brett and Sutherland 1965; Evans 1984). (b) Effects of water temperature and stock on the baseline oxygen consumption rates of largemouth bass. Closed squares represent central Illinois pure stock (IL), closed triangles represent southeastern Wisconsin stock (WI), closed circles represent WI × IL stock and open squares represent IL × WI stock (see Cooke et al . 2001a, for details). (c) Effects of ambient oxygen concentrations on the baseline metabolic rates in largemouth bass at 20◦ C (closed circles), 25◦ C (closed triangles), and 30◦ C (closed squares). (Redrawn from Cech et al . 1979. American Fisheries Society.)
and white crappie (Pomoxis annularis) exist. The published results for critical swimming speeds show that considerable variation exists within the Centrarchidae (see Table 8.3). For example, values for UCrit typically range from between 2 and 3.5 BL s−1 for smallmouth bass (McDonald et al . 1991; Cooke and Bunt 2001; Peake 2004, but see Larimore and Duever 1968 for work on juveniles), about 3 BL s−1 for bluegill (Kelsch 1996), 2 BL s−1 for white crappie (Parsons and Smiley 2003), 3 BL s−1 for pumpkinseeds (Brett and Sutherland 1965) and as high as 3.5–8 BL s−1 for largemouth bass (Dahlberg et al . 1968; Hocutt 1973; Farlinger and Beamish 1977; Farlinger and Beamish 1978, but see Ostrand et al .
Physiology and organismal performance of centrarchids
215
Table 8.2 Maximum muscle lactate concentrations (umol g−1 wet tissue) following exhaustive exercise stress in various centrarchid and noncentrarchid species of fish. Species
Muscle lactate (umol g−1 wet tissue)
Largemouth bass (adult) Smallmouth bass (adult) Rainbow trout (adult)
Source
∼20
Kieffer et al. 1996
∼15
Suski et al. 2004
∼18
Kieffer et al. 1995a
∼30
Peake and Farrell 2004
41
Schulte et al. 1992
∼30
Kieffer et al. 1994
Coho salmon
∼45
Farrell et al. 2001b
Atlantic salmon (adult)
∼45
Wilkie et al. 1996
Brook charr (adult)
∼33
Kieffer et al. 1996
Sea lamprey (adult)
∼25
Boutilier et al. 1993
Yellow perch
∼24
Schwalme and Mackay 1991
Atlantic sturgeon (juvenile)
∼6
Kieffer et al. 2001
a indicates post-angling lactate levels. b indicates lactate levels following capture by commercial fishers.
2005, for exceptions on juveniles). Critical swimming speeds of centrarchids are similar to those for other freshwater and marine fish species, including: brown trout (UCrit ∼1.8; Altimiras et al . 2002), rainbow trout (UCrit ∼3.0; Kieffer et al . 1998; UCrit ∼1.5; Jain and Farrell 2003), common snook (Centropomus undecimalis; UCrit ∼2.2–8.9 BL s−1 , depending on size; Tolley and Torres 2002), cut-throat trout (Oncorhynchus clarki clarki ; UCrit ∼4; MacNutt et al . 2004), coho salmon (UCrit 1.5–2.5; Lee et al . 2003a), and walleye (Sander vitreus; UCrit ∼0.8–1.2; Peake et al . 2000). Although differences in UCrit values between centrarchid species may reflect differences in the methodology(ies) used to determine critical swimming speeds (e.g., size of fish, time increments for testing; Farlinger and Beamish 1977; Peake et al . 2000), it is highly possible that the variability in these values reflects various abiotic and biotic factors (see Section 8.3.4) or differences in life history and ecology of the various centrarchid species (see Chapter 13).
8.3.4 Effects of abiotic factors on UCrit 8.3.4.1 Temperature effects The effects of temperature on swimming performance in fish are well known (Brett 1964; Hocutt 1973; Randall and Brauner 1991; Keen and Farrell 1994; Kieffer et al . 1998; Myrick and Cech 2000; Cooke et al . 2001a; Lee et al . 2003a,b; MacNutt et al . 2004; O’Steen and Bennett 2003). In general, increases in temperature to an optimum improve swimming performance by enhancing biochemical rates (Franklin 1998), skeletal muscle contractility (Rome et al . 1990), and cardiac performance (Kolok and Farrell 1994a,b). In an early study by Larimore and Duever (1968), groups of smallmouth bass fry were acclimated to water temperatures ranging from 5 to 35◦ C. The maximum swimming speed for fish acclimated to a particular temperature increased with successively higher test temperatures until 30◦ C (see Figure 8.2), with fry acclimated to 30◦ C achieving a maximum swimming speed twice that of fry acclimated to 5◦ C. These authors also acutely challenged acclimated animals to different temperatures (i.e. 5◦ C acclimated fish tested at 10, 15, and 20◦ C) and found that critical swimming speeds also increased with increases in temperature, but the range of performance changed in comparison in the response of acclimated fish. For example, only fish acclimated to 5 or 10◦ C performed well at 5◦ C and only fish conditioned to 35◦ C performed well at 35◦ C. Hocutt (1973) measured the swimming performance of largemouth bass exposed to a rapid temperature change (temperature range: 15–35◦ C) and also noted positive correlations between increasing temperature and swimming performance.
216
Centrarchid fishes
Table 8.3 Critical swimming (UCrit) values for various species of centrarchids. Species
Size (length/mass)
Temperature (◦ C)
UCrit (cm s−1 )
UCrit (Bl s−1 )
Source
13.5–17.5 cm
13
–
∼2.4
Kelsch 1996
13.5–17.5 cm
25
–
∼3
Kelsch 1996
13.5–17.5 cm
30
–
∼2.8
Kelsch 1996
Pumpkinseed
12.7 cm
20
–
3.01
Brett and Sutherland 1965
White crappie
16.5–17.5 cm
25
34.7
–
8.01 cm
5
6.16
0.76
Smiley and Parsons 1997c
7.63 cm
15
13.59
1.78
Smiley and Parsons 1997c
8.00 cm
25
13.86
1.73
Smiley and Parsons 1997c
Bluegill
Largemouth bass
Parsons and Sylvester 1992
5–10 cm
5–7
∼10
–
Parsons and Smiley 2003
5–10 cm
15–18
∼16
–
Parsons and Smiley 2003
5–10 cm
24–27
∼20
–
Parsons and Smiley 2003
10–15 cm
5–7
∼11
–
Parsons and Smiley 2003
10–15 cm
15–18
∼20
–
Parsons and Smiley 2003
10–15 cm
24–27
∼28
–
Parsons and Smiley 2003
15–20 cm
5–7
∼10
–
Parsons and Smiley 2003
15–20 cm
15–18
∼15
–
Parsons and Smiley 2003
15–20 cm
24–27
∼30
–
Parsons and Smiley 2003
20–25 cm
5–7
∼15
–
Parsons and Smiley 2003
20–25 cm
15–18
∼20
–
Parsons and Smiley 2003
20–25 cm
24–27
∼30
–
Parsons and Smiley 2003
25–30 cm
5–7
∼15
–
Parsons and Smiley 2003
25–30 cm
15–18
∼20
–
Parsons and Smiley 2003
25–30 cm
24–27
∼25
–
Parsons and Smiley 2003
9.3–12.8 cm 9.3–12.8 cm
5 5
– –
∼2.2 ∼1.7
Kolok 1991d Kolok 1991e
9.3–12.8 cm
5
–
∼1.5
Kolok 1991f
9.3–12.8 cm
10
–
∼2.9
Kolok 1991g
9.3–12.8 cm
10
–
∼2.8
Kolok 1991h
9.3–12.8 cm
10
–
∼2.3
Kolok 1991i
10.1 cm
5
20
1.98
Kolok 1992
9.7 cm
20
35.7
3.68
Kolok 1992
15.9 cm
6
30.1
1.90
Cooke et al. 2001a
17.0 cm
12
34.6
2.06
Cooke et al. 2001a
16.6 cm
18
35.2
2.01
Cooke et al. 2001a
8.6 cm
20
∼60–70
∼7.2–8.4
Ostrand et al. 2004a
12.2 cm
25
41.63
3.41
10.4 cm
25
–
∼3.5
Farlinger and Beamish 1978 Farlinger and Beamish 1978
5.2–6.4 cm
30
29.81
8.08
Hocutt 1973 (continued)
Physiology and organismal performance of centrarchids
217
Table 8.3 (continued). Species Smallmouth bass
Size (length/mass)
Temperature (◦ C)
UCrit (cm s−1 )
UCrit (Bl s−1 )
2.2 cm 2.2 cm
5 10
– –
2.2 4.7
Larimore and Duever 1968b Larimore and Duever 1968b
2.2 cm
15
–
6.8
Larimore and Duever 1968b
2.2 cm
20
–
10.2
Larimore and Duever 1968b
2.2 cm
25
–
11.8
Larimore and Duever 1968b
2.2 cm
30
–
13.6
Larimore and Duever 1968b
2.2 cm
35
–
11.5
Larimore and Duever 1968b
Source
10.1 g
16
–
2.0
McDonald et al. 1991
∼30 cm
–
83.75
2.79
S. Peake, unpublished data
31.0 cm
17
111.3
3.59
26.2–37.8 cm
15–20
50–118
–
Bunt et al. 1999
42.1–48.9 cm
20–24
47–69
–
Bunt 1999
Cooke and Bunt 2001
a Values depend on diet. See Ostrand et al. (2005), for details. b Fry were swum in groups of 20. c Fish held under 12:12 (L:D) photoperiod. See Smiley and Parsons (1997) for additional details and UCrit values. d Winter acclimatized (5◦ C) fish. ◦ e
Laboratory acclimated to 5 C, photoperiod 9:15 L:D.
f Laboratory acclimated to 5◦ C, photoperiod 12:12 L:D. g Spring acclimatized (10◦ C) fish. h Laboratory acclimated to 10◦ C, photoperiod 9:15 L:D. ◦ i
Laboratory acclimated to 10 C, photoperiod 12:12 L:D.
Smallmouth bass
16 UCrit (cm s −1)
14 12 10 8 6 4 2 0 5
10
15
20
25
30
35
Temperature (°C) Figure 8.2 Mean critical swimming speed of groups of smallmouth bass fry acclimated to various temperatures (data from Larimore and Duever 1968. American Fisheries Society).
Parsons and Smiley (2003) measured critical swimming speeds in white crappie that were seasonally acclimatized to winter (5–7◦ C), spring (15–18◦ C), and summer (24–27◦ C) temperatures. These authors noted the typical pattern that UCrit values increased in fish at the higher temperatures (see earlier and Figure 8.2). They also documented that the highest number of nonperforming fish (those with UCrits 90% mortality for some centrarchids—although this is typically during competitive angling events at high water temperatures). There are a number of strategies that can be employed to minimize hooking mortality of centrarchids. For example, barbless hooks have been shown to reduce tissue damage, bleeding, and air exposure duration (through more rapid hook removal) in rock bass (Cooke et al . 2001b). In addition, use of artificial baits rather than organic/live baits tends to result in shallower hooking and less injury and mortality, as has been observed for bluegill (Siewert and Cave 1990) and smallmouth bass (Clapp and Clark 1989). Interestingly, however, scented lures do not appear to cause greater injury or mortality than nonscented lures for smallmouth bass (Dunmall et al . 2001). Hook type also has the potential to influence hooking depth.
344
Centrarchid fishes
For example, circle hooks have been shown to reduce deep hooking and instead facilitate jaw hooking in a number of fish species (Cooke and Suski 2004). However, there is little evidence that circle hooks make a meaningful difference for centrarchids including rock bass (Cooke et al . 2003a), largemouth bass (Cooke et al . 2003c), bluegill (Cooke et al . 2003e), or pumpkinseed (Cooke et al . 2003e). One of the primary reasons for inconsistent results with circle hooks in centrarchids is the wide range of mouth morphologies and body sizes that makes it difficult to select an appropriately sized circle hook (Cooke et al . 2005a). Research into the sublethal stressors that result from a catch-and-release angling event is also on the rise. Various components of a catch-and-release angling event have been determined to be quite stressful and harmful to the angled fish. An angling event for a fish of any species is essentially a period of intense aerobic and anaerobic exercise that can lead to a host of physiological changes such as depletion of energy stores, increases in lactate in muscle, and disturbances to acid/base balance and osmoregulation. As angling duration increases, the magnitude of these effects also increases. In largemouth bass, the severity of physiological disturbances such as increased levels of lactate, chloride, and plasma glucose has been shown to correlate positively with angling duration (Gustaveson et al . 1991). Similarly, Kieffer et al . (1995) found that physiological disturbance in white muscle (increased lactate and metabolic protons) as well as energy depletion were also more severe as angling duration increased. Smallmouth bass that were angled to exhaustion were also found to require longer periods of time to recover from the cardiac disturbance caused by angling (Schreer et al . 2001). As such, efforts to minimize angling duration (such as use of appropriate equipment) are helpful in minimizing anaerobic physiological disturbance. Air exposure has also been proven to cause negative physiological disturbances for a variety of fish species, and the results of these studies are applicable to centrarchid fishes. While exposed to air, gill filaments can adhere to each other as a result of the collapse of gill lamellae (Boutilier 1990). During this time, the fish are essentially deprived of oxygen and subjected to induced anoxia that can reduce the amount of oxygen delivered to tissues (Ferguson and Tufts 1992). Air exposure has also been shown to increase the severity of various physiological disturbances associated with angling (Ferguson and Tufts 1992). Air exposure also induces negative cardiovascular disturbances in both rock bass and smallmouth bass, with individuals exposed to air for longer durations requiring longer to return to basal levels (Cooke et al . 2001b; Cooke et al . 2002). In addition, if fish are held in nets during air exposure, scale loss can occur, which has been identified as a precursor to mortality in bluegill (Barthel et al . 2003). Simply minimizing air exposure duration, or ideally eliminating it, is the best way to minimize the negative consequences of air exposure. Fish should be held by wet hand rather than in nets if possible. Mortality among angled individuals has also been linked to water temperature during the angling event. Physiological disturbance and associated mortality have been shown to occur more often when fish are angled at extremely high temperatures. Fish subjected to exhaustive exercise show increased levels of physiological disturbance (lactate and metabolic protons in the blood) at higher water temperatures. High water temperatures have been found to exacerbate the physiological disturbances associated with angling duration (Meka and McCormick 2005). In smallmouth bass and largemouth bass, the recovery from the cardiac disturbance associated with angling also varies with water temperatures (Schreer et al . 2001; Cooke et al . 2003d). Also, mortality of angled largemouth bass increased with water temperatures, particularly beyond a threshold in the high 20◦ Cs (Wilde 1998). Anglers should avoid fishing at high water temperatures (i.e. >30◦ C) or if that is not possible, minimize other stressors such as air exposure and angling duration. One of the only studies evaluating the interaction between multiple stressors revealed that mortality rates of bluegill were highest when warm water temperatures (>26◦ C) were coupled with prolonged air exposure (>120 s; Gingerich et al . 2007). Mortality was negligible when air exposure was eliminated even though water temperatures were high. Centrarchid fishes are particularly vulnerable to angling during their reproductive period. Briefly, during the reproductive period male centrarchids construct nests, court females, and provide vigorous defense to the developing brood for some amount of time ranging from a few days to a month or longer (Cooke et al . 2006). Males angled prior to the reproductive period were found to produce fewer and smaller offspring than nonangled males (Ostrand et al . 2004). During the reproductive period, when the male is removed from the nest by anglers, predators can devour the brood, thereby directly reducing the fitness of the individual (Neves 1975; Philipp et al . 1997; Steinhart et al . 2004a; Hanson et al . 2007). Research has demonstrated that brood loss to predation is responsible for nest abandonment in angled centrarchids, rather than physiological disturbances associated with the catch-and-release angling event (Suski et al . 2003b). As the time the male is kept away from the nest increases, the percentage of the brood consumed by predators also increases, increasing the likelihood of abandonment by the male (Kieffer et al . 1995; Philipp et al . 1997). More importantly, the individual males
Contemporary issues in centrarchid conservation and management
345
that are most vulnerable to angling are often largest and most aggressive fish that have nests with the highest numbers of eggs (Suski and Philipp 2004). Because males with the largest broods (and therefore the largest potential to contribute to year class formation) are the most likely to be caught by anglers, negative population-level effects may occur if nesting centrarchids are subjected to angling pressure during the nest-guarding period (Suski and Philipp 2004). Ridgway and Shuter (1996) predicted that as the number of angled male smallmouth bass increased across the reproductive period, the abundance of age-0 fish would decrease due to brood predation and nest abandonment. The negative effects of a catch-and-release event linger with angled male even after he returns to the nest and resumes nest-guarding behavior. Previously angled largemouth bass have reduced locomotory potential when compared to nonangled individuals while guarding nests (Cooke et al . 2000). This disturbance coupled with brood loss due to predation during the angling event could increase nest abandonment and loss of fitness (Cooke et al . 2002). Also, during the parental care period, nestguarding male bass are not free to forage normally and are engaged in a set of energetically costly behaviors (Hinch and Collins 1991; Cooke et al . 2006). Physiological disturbance during this time period of high stress, low food consumption, and high energetic demands may be particularly debilitating to the ability of the male to successfully raise a brood (Cooke et al . 2002). Based on this information, it appears prudent that angling should be restricted during this period. Interestingly, there are very few jurisdictions where angling during bass reproductive period is actually prohibited (e.g., Ontario; Quinn 1989). The topic of restrictive angling regulations during centrarchid spawning is a contentious topic in the angling and fisheries management communities with several weak arguments supporting the more liberal perspective: (i) Work to date has focused on the northern populations where the growing season is much shorter. As such, the findings from the “north” do not apply elsewhere. Although it is quite likely that there are latitudinal differences in reproductive biology and growing season, there is still no evidence to support the idea that angling during the reproductive period is either beneficial or benign. (ii) If catch-and-release angling for nesting bass is harmful, then why is there not evidence of massive collapse in intensive fisheries? This argument is difficult to address given the challenge of monitoring fish populations, particularly adult centrarchids. However, in many jurisdictions where angling for nesting fish is permitted, there are also supplemental stocking programs that could mask negative effects. What is clear is that there is a need for additional research to evaluate the link between reproductive success and recruitment, as well as comparative research in warmer clines (than Ontario) to test the hypothesis that angling during the reproductive period is not an issue for black bass in the south. Because of the contentiousness of this issue, only with additional data in hand from more regions will managers be able to make informed decisions based on scientific fact.
12.2.1.9 Competitive angling events An area of specific concern for centrarchid conservation is the growing popularity and prevalence of angling tournaments targeting black basses, specifically largemouth and smallmouth bass (Shupp 1979; Schramm 1991). During tournaments, angled fish face multiple situations that can result in physiological disturbances, and that may ultimately lead to mortality (Suski et al . 2003a; Siepker et al . 2007). That said, competitive angling events provide unique opportunities to assess black bass fisheries through tournament monitoring programs, and recent estimates of mortality rates confirm that, at many tournaments, most fish are returned to the water alive following the completion of the angling day (Wilde 1998). In some cases, tournament-collected data provide the only information on difficult-to-sample water bodies such as medium-sized rivers (e.g., Cooke et al . 1998). During competitive angling events, multiple fish are often confined to livewells for extended periods of time. While retained in a livewell, fish can quickly face extremely poor water quality as ammonia and dissolved carbon dioxide levels rise and dissolved oxygen decreases (Hartley and Moring 1993; Kwak and Henry 1995), resulting in a state of hypoxia and a host of physiological disturbances associated with air exposure (Furimsky et al . 2003). These conditions are exacerbated by high densities of fish confined within the livewell (Cooke et al . 2002). Also, when confined at high densities in a livewell, fish show increased physiological and cardiac disturbances (Cooke et al . 2002). Another source of mortality during livewell confinement may be disturbances associated with wave conditions (Kwak and Henry 1995). Recent video analysis of largemouth bass behavior during livewell confinement found that during low level disturbances, individuals remained active (Suski et al . 2005). During high intensity disturbance, largemouth bass tended to face the direction of the disturbance while staying near the bottom of the livewell, allowing them to avoid repeatedly hitting
346
Centrarchid fishes
the sides of the livewell (Suski et al . 2005). Anglers should ensure that livewells are properly circulated so as keep water quality favorable (Gilliland 2002), keep densities of fish in livewells as low as possible (Cooke et al . 2002), and operate their boat in a manner that minimizes disturbance associated with wave action (Suski et al . 2005). The addition of livewell conditioners has also been suggested as a way to help fish recover from livewell confinement. Currently, there is conflicting evidence as to whether livewell conditioners enhance recovery, and more research needs to be conducted on this topic to conclusively prove the benefit or harm of livewell conditioners (Gilliland 2002; Cooke et al . 2002). Recent research revealed that hyperoxygenation or decreasing livewell water temperatures did not aid in recovery of bass during livewell retention and actually increased physiological disturbance (Suski et al . 2006). Livewells are always preferable to stringers or fish baskets for temporarily retaining centrarchids (Cooke and Hogle 2000). During an angling tournament, fish are subjected to repeated air exposure. While angling, fish are often air exposed as anglers repeatedly place and remove them from livewells while culling individuals from the final catch (Cooke et al . 2002). At the end of a tournament, fish are subjected to weigh-in processes before release. Often, these processes include extended periods of confinement in nonaerated water in holding areas followed by air exposure during the weighing (Suski et al . 2004). During this time period, bass showed significant metabolic disturbances related to air exposure including increases in tissue lactate, cardiac disturbances, and decreases in tissue energy stores (Suski et al . 2004). The cumulative effect of these stressors may lead to later mortality by impairing the ability of released individuals to survive (Cooke et al . 2002). To minimize the physiological disturbances of weigh-in, it has been suggested that tournaments minimize air exposure throughout, but especially during weighing by keeping fish in well-aerated water as often as possible (Furimsky et al . 2003; Suski et al . 2004). Recent innovations have included the development of a water weigh-in system that almost eliminates air exposure (Suski et al . 2004). In addition to the disturbances that bass can experience during an angling tournament, research has shown that events following the release of fish can have potential for population-level impacts. Specifically, high concentrations of fish are often released in a small area at the conclusion of an angling tournament resulting in a “stockpiling” of fish that can persist for short periods of time. Ridgway (2002), for example, reported that over 2 weeks had passed before translocated largemouth bass moved 400 m from a common release site, whereas Wilde and Paulson (2003) showed that 63% of tournament-caught largemouth bass remained within 0.5 km of their release site following a 43-day monitoring period. Ridgway and Shuter (1996) showed that displaced smallmouth bass remained near their release site from 1 m apart) tend to nest near simple cover (e.g., bases of logs, rocks, or macrophytes) and defend a territory exceeding the nest perimeter (>2.5 m, Colgan and Ealey 1973; Avila 1976; Winemiller and Taylor 1982; Colgan and Brown 1988; Ridgway 1988; Jennings and Philipp 1992b; Scott 1996). Colonies of nests, consisting of several to hundreds of abutting nests, tend to occur in shallow open water, and in dense colonies nest defense is constrained primarily to the nest perimeter (Hunter 1963; Colgan et al . 1981; Gross and MacMillan 1981; Gross 1982). Spawning can occur immediately after nest construction or be delayed for several days, during which the male defends the nest and surrounding territory and waits for spawning-ready females (Carr 1946; Kramer and Smith 1962; Boyer and Vogele 1971; Miller and Kramer 1971; Avila 1976; Vogele 1975a; Colgan and Gross 1977; Gross and Nowell 1980; Cooke et al . 2001b). Male aggression intensifies during the courtship and spawning period. Males over nests display to nearby or approaching males and females using combinations of many behaviors (e.g., caudal sweeping, nest hovering, fin spreading, mouth gapes, jaw snaps, lateral displays, substrate biting, and opercular spreads). Male to male aggressive interactions, including combat, are not uncommon, particularly among colonial-nesting species. Males most frequently rush toward an interloper with a quick retreat to the nest (thrust, Miller 1963), but if the intruder does not retreat, males laterally display, spread opercles, or actually ram, push, bite, or jaw grasp the other male. Much of male aggression is directed at or near the head and opercular area, but frayed fins and body abrasions of males attest to the vigorousness of male aggression in defense of the nesting territory (Hunter 1963; Keenleyside 1967, 1971; Colgan and Gross 1977; Gross and Nowell 1980). Male courtship of females may be preceded by attempts to repulse females near the nest, behaviors that coax or guide the female to the nest, or both. Repeated repulsion of approaching females by males is documented in Archoplites (Mathews 1965), Ambloplites (Gross and Nowell 1980; Petrimoulx 1984; Noltie and Keenleyside 1987b), Lepomis (e.g., Hunter 1963; Huck and Gunning 1967; Keenleyside 1967; Ballantyne and Colgan 1978a,b,c), and Pomoxis (Siefert 1968). If ready to spawn, a female, assuming a subordinate demeanor, continues to slowly approach the nest despite repeated attacks by the male. Male-leading or -guiding courtship behaviors are known in Lepomis, Micropterus, and Centrarchus, although Lepomis females often enter nests with little or no overt courtship (Carr 1942; Dickson 1949; Hunter 1963; Keenleyside 1967; Chew 1974; Coble 1975; Vogele 1975a; Avila 1976; Gross 1982; Ridgway et al . 1989; Lukas and Orth 1993; Cooke et al . 2001b). Repulsing or guiding male behaviors directed at females may be species or context specific, are difficult to separate cleanly into courtship or aggression, and often co-occur (Keenleyside 1967; Ballantyne and Colgan 1978a,b,c). Once a pair is situated over the nest, they orient broadside and head to head and swim in slow, tight circles over the nest. The pair settles to the substrate, and egg deposition occurs as the female tilts away from the male and presses her vent near the substrate; the male presses his vent to the female’s while remaining upright or rolling toward the female. Egg and sperm release is accompanied by shuddering in both sexes; the demersal, adhesive eggs adhere to the nest substrate and to one another in clumps. Typically the pair rests, then repeats the sequence multiple times, until the male chases the female out of the nest. Rests between spawning bouts tend to shorten as the spawning event continues. These sequences may be in quick succession if the pair is not interrupted by intruders, but completion of spawning with a single female may occur over extended periods (15 minutes to 3.5 hours), even without interruption (Siefert 1968; Neves 1975; Vogele 1975a; Gross 1982, 1991; Isaac et al . 1998; Cooke et al . 2001b). After spawning, males aggressively guard the eggs and larvae, but the length of male parental care after the eggs hatch differs among genera and species within genera. Today, centrarchids are the primary focus of the recreational fishing industry in the United States and much of southeastern Canada. The relatively large size of many centrarchids, vulnerability to natural baits or artificial lures, and the excellent taste of the flesh combine to create a popular sport fishery worth billions of dollars a year. The black basses (Micropterus), particularly the Florida bass and largemouth bass, the bream or panfishes (Lepomis), especially the bluegill, and the crappies (Pomoxis) are sought by anglers more than any fresh or saltwater sport fishes in the United States. Angler numbers and days spent fishing for centrarchids dwarf those reported for salmonids, walleye, or saltwater fishes (USFWS 2002). A prodigious body of information is available on centrarchid natural history. Most research, however, has focused on a relatively few but important sport fish species, and there is no single-source recent summary of natural history information for all species in family. The objective here is to provide synopses of the characteristics and the natural history of the
Centrarchid identification and natural history
377
8 genera and 34 species of centrarchid fishes and to provide a dichotomous key to the family. A secondary objective of this chapter is to highlight species for which information on their natural history is lacking, fragmentary or anecdotal.
13.2 Generic and species accounts The bulk of the chapter consists of a separate account for each genus and each species within a genus, with the exception of monotypic genera. Only species accounts are given for monotypic genera. Within the characteristics sections of generic and species accounts, the definition of counts, standard length (SL), total length (TL), and other measurements follow standard ichthyological methods (see Page and Burr 1991; Jenkins and Burkhead 1994; Boschung and Mayden 2004) or are given in the citations associated with that section. Counts are presented as a total range, that is, 19 to 25; a modal (usual) count followed by a range, that is, usually 22, 19 to 25; or the most frequently encountered range of counts (ca. ≥90%) and the extremes, that is, (19)21 to 23(25). Only published sources were used to designate a confirmed freshwater mussel host (e.g., mussel larvae successfully infected and transformed on a centrarchid host). A putative host is similarly defined, except that the data are from unpublished sources and need verification. Published or unpublished accounts of mussel larvae infection on a centrarchid species without observation of transformation to the juvenile stage are not included.
13.3 Acantharchus pomotis (Baird) 13.3.0.1 Mud sunfish Characteristics: Moderately oblong and robust body, depth 105 mm TL (Pardue 1993). Reproduction: Maturity is reached at age 1+ and a minimum size of 66 to 140 mm TL. Spent females, egg sizes, and gonad to body weight ratios suggest that the mud sunfish begins and completes spawning at temperatures as low as 7 to 10◦ C (Pardue 1993), which is lower than minima reported for other centrarchids. The spawning period apparently extends from December to May in North Carolina and into June in New Jersey at water temperatures of 7 to 20◦ C (Breder 1936; Pardue 1993). The ovaries enlarge in the early fall and continue developing over winter (Pardue 1993), which is likely an adaptation for early spawning. Reproductive behaviors are essentially unknown. Males have been observed or captured over small depressional nests near the shoreline of lakes or near the banks of headwater streams in water 15 to 30 cm deep (Fowler 1923; Marcy et al . 2005). Mud sunfish produce audible grunting noises (Gerald 1971), but linkage with reproduction is undocumented. Mature ovarian eggs range from 0.7 to 1.1 mm diameter (Pardue 1993). At a median size of 128 mm TL, a female can produce 2304 mature eggs (range: 1515 at 114 mm TL to 3812 at 144 mm TL; data from Pardue 1993), which is one of the lowest batch fecundities among centrarchids (see also Ambloplites and Enneacanthus). Female allocation of energy to reproduction is also low relative to most centrarchids with peak female gonad to somatic weight values of 3% (Pardue 1993). Mature ovarian egg size is similar to that in Lepomis and may indicate a similar duration of male care provided to the embryos and larvae (Gross and Sargent 1985), but the combination of low batch fecundity and low female energy allocated to reproduction differs from reproductive patterns observed in all other centrarchids. Nest associates: None known. Freshwater mussel host: None known. Conservation status: The mud sunfish is widely distributed but not common anywhere. The species appears to be secure where its lowland habitats are undisturbed, particularly in the central portions of its Atlantic Coastal Plain range (North and South Carolina). Populations to the north and south are considered possibly extirpated (New York), imperiled (Delaware and Maryland), or vulnerable (Virginia, Georgia, and Florida) (NatureServe 2006). Similar species: All other centrarchids have ctenoid scales (cycloid in Acantharchus), and except for Enneacanthus, deeply to shallowly emarginate caudal fins (rounded in Acantharchus and Enneacanthus). Enneacanthus possess three anal fin spines (4–6 in Acantharchus). Systematic notes: The phylogenetic relationships of the monotypic genus Acantharchus to other centrarchid genera is the least resolved within the family. Phylogenetic analyses place the species as sister to all other centrarchids or as resolved within a clade of all centrarchid genera but Lepomis and Micropterus (Roe et al . 2002; Near et al . 2004, 2005). The species shows evidence of polytypy. A subspecies described from the Okefenokee Swamp region (Suwannee River drainage, Georgia) as A. pomotis mizelli (Fowler 1945) was based on little comparative data. In an extensive study of geographic variation, several meristic characters of populations in eastern Gulf of Mexico drainages diverged significantly from those of populations in Atlantic Slope drainages. Multivariate analyses of morphological characters suggested that a contact zone between northern Atlantic Slope populations and Gulf Slope populations exists in Atlantic Slope drainages
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of Georgia and Florida (Cashner et al . 1989). Resolution of the evolutionary distinctiveness of the two geographic groups awaits molecular phylogeographic analysis. Importance to humans: The mud sunfish is one of the least known of all centrarchids, even to avid sport fishers, fisheries biologists, and most ichthyologists. The species is apparently rarely taken by hook and line and can go uncaught and unnoticed by anglers even when it occurs in heavily fished ponds (Mansueti and Elser 1953). Unfortunately, so little is known about the species that its ecological function and value in lowland stream and wetland ecosystems cannot be evaluated, but its adaptability to such habitats and distribution across a broad latitudinal band suggest a long evolutionary history in those environments and a potentially important functional role. The basal phylogenetic relationship of Acantharchus within the centrarchids may provide an important key for unraveling the relationship of the centrarchids to other percoid fishes, a relationship that is currently unknown. Likewise, study of its reproductive biology and behavior could illuminate the evolutionary history of complex reproductive strategies and associated behaviors observed in other centrarchids.
13.4 Ambloplites Rafinesque The monophyletic genus Ambloplites, often referred to collectively as rock basses, is endemic to eastern North America and contains four species consisting of two sister group pairs: Ambloplites ariommus (shadow bass) and Ambloplites rupestris (rock bass) form one sister pair and Ambloplites cavifrons (Roanoke bass) and Ambloplites constellatus (Ozark bass), the other. Ambloplites is sister to the monotypic genus Archoplites, represented by the Sacramento perch, and these two genera are sister to the genus Pomoxis (Near et al . 2004, 2005). The genus is distributed broadly across eastern North America, mostly east of the Great Plains, from southern Canada to the Gulf Coastal Plain, but the natural ranges of all four species are allopatric within this region. The Roanoke bass–Ozark bass sister pair occupies some of the smallest ranges of any North American sport fish. The Roanoke bass is endemic to Atlantic Coast drainages of Virginia and North Carolina and the Ozark bass mostly to the White River of Arkansas and Missouri. The range of the shadow bass is essentially disjunct; part of the range includes drainages of the eastern Gulf Slope and lower Mississippi River and the remainder includes drainages of the Ouachita Mountains, Arkansas River Valley, and Ozark Plateau. The rock bass, the most broadly distributed member of the genus, has been introduced and is widely established outside its native range in both eastern and western North America (Cashner and Suttkus 1977; Fuller et al . 1999). Intentional (or suspected) introductions of rock bass and other species of Ambloplites into the ranges of congeners has obscured natural ranges, has produced introgressed populations, and threatens the genetic integrity of species within the genus, particularly the range-restricted endemics (Cashner and Suttkus 1977; Cashner and Jenkins 1982; Jenkins and Burkhead 1994; Koppelman et al . 2000). Ambloplites appear to differ from most other centrarchids, except their sister genus Pomoxis, in several aspects of reproductive behavior, but detailed, multiple observations are available only for rock bass. Male Ambloplites apparently do not use caudal sweeping to clear nesting areas as is common in most other centrarchid males (Miller 1963). Ambloplites males use a combination of behaviors to construct the nest, including undulations of the anal fin, sweeping of the pectoral fins, and pushing material forward with outstretched pectoral fins (bulldozing, Gross and Nowell 1980; Petrimoulx 1984; Noltie and Keenleyside 1987b). Males orient slightly head downward and use alternating strokes of the pectoral fins for fanning the eggs, similar to Pomoxis, rather than the horizontally oriented and primarily caudal- fin fanning as described for Lepomis or Micropterus (Carr 1942; Miller 1963; Gross and Nowell 1980; Noltie and Keenleyside 1987b). Males show no overt courtship of females, and mate choice appears to be restricted to male acceptance of females (Gross and Nowell 1980; Petrimoulx 1984). Males aggressively and persistently repel and even attack females approaching the nest, spawning only with the most persistent, submissive females, behaviors in contrast to the active leading or guiding behaviors of nest-defending males toward females in other genera (e.g., Lepomis and Micropterus). The relative position of the male to the female during spawning also appears to differ in, and perhaps among, Ambloplites. The male of the Roanoke and Ozark bass occupies a central nest position during pairings with females rather than a position outside the female (toward the nest rim); the rock bass male takes an outside nest position in spawning if circling occurs, but occupies a central position when no nest circling occurs (Gross and Nowell 1980; Petrimoulx 1984; Noltie and Keenleyside 1987b; Walters et al . 2000). Members of Ambloplites are popular sport and food fishes and are commonly taken by anglers. In Missouri, three species, the shadow bass, rock bass, and Ozark bass, comprise 10% of the catch and harvest of fishes in streams (Koppelman
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et al . 2000). Many individuals are caught incidentally with the same lures and tackle used by anglers seeking smallmouth, spotted, and redeye basses, which frequently co-occur with species of Ambloplites. Anglers specifically seeking rock basses use small lures and spinners, lures imitating minnows, or live bait, particularly dobsonfly larvae (hellgrammites) and small crayfishes (Nielsen and Orth 1988; Ross 2001). Anglers often refer to these fishes as “redeyes” because of the conspicuous red pigment in their iris or “goggle eyes” because of their relatively large and conspicuous eyes (Etnier and Starnes 1993; Koppelman et al . 2000). Generic characteristics: Moderately compressed, elongate body, depth 0.42 of SL. Eye large, diameter typically >0.30 of head length. The pattern of dark blotches alternating with lighter areas on body in young is retained in adults, so that adults and young resemble the appearance of young A. rupestris. Preopercle sharply serrate to weakly crenate to entire at the angle. Dorsal fin elements, (20)22 to 23(24); anal fin elements, (15)16 or 17(18). Cheeks fully scaled with large, exposed scales. Cheek scale rows, (5)6 or 7(8); lateral line scales, (34)38 to 43(45); scale rows above lateral line, (5)6 or 7(8); scale rows below lateral line, (11)13 to 15(16); diagonal scale rows, (18)22 or 23(24); and breast scale rows, (13)16 to 18(20). One circular patch of teeth on tongue (Cashner 1974; Cashner and Suttkus 1977; Page and Burr 1991). Size and age: Typically reach 40 to 120 mm TL at age 1. Large individuals measure 160 to 203 mm TL, rarely exceed 340 g, and reach age 6+ to 9+ (maximum 220 mm TL); Missouri and Arkansas populations can apparently reach larger sizes (at least 254 mm TL) than other populations (Viosca 1936; Robison and Buchanan 1984; Page and Burr 1991; Pflieger 1997; C. S. Schieble, University of New Orleans, personal communication). World angling record, 820 g, Arkansas (IGFA 2006). Females may outlive males, and males slightly exceed females in average maximum size and weight, but growth curves for the sexes are similar (C. S. Schieble, University of New Orleans, personal communication). Coloration: Light green to brown on sides with irregular marbling of brown or gray dark blotches alternating with lighter areas, blotches often joined dorsally to form saddles. Scales on sides bear a dark, triangular spot at the base (apex forward), producing a pattern of longitudinal lines that run through but are often obscured by the light and dark pigmented areas. Lower sides and belly transitioning to straw color (Viosca 1936; Cashner 1974; Page and Burr 1991). Large breeding males have a distinct darkening of the membranes in the pelvic and anal fins from the fin tips to the base and distinct black, threadlike filaments on their pelvic fins. These filaments are yellow to white in females (C. S. Schieble, University of New Orleans, personal communication).
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Native range: The range of the shadow bass is disjunct. The species occupies Gulf Slope drainages from the Apalachicola River west to the lower Mississippi River, including the Mobile Basin, and also occurs in the Red, Ouachita, Arkansas, St. Francis, and Black rivers (Page and Burr 1991). Habitat: The shadow bass inhabits gravel, sand, and mud-bottomed creeks and small to medium rivers with low levels of turbidity and sedimentation. The species is almost always associated with pools and cover of boulders, logs, log complexes, or rootwads; water willow or other aquatic vegetation in shallow water often harbors young-of-the-year (Probst et al . 1984; McClendon and Rabeni 1987; Page and Burr 1991; Pflieger 1997, reported as rock bass; C. S. Schieble, University of New Orleans, personal communication). In a large-scale tagging study (Funk 1957), shadow bass (reported as rock bass) were regarded as sedentary, but 48% and 31% of recaptured individuals moved at least 1.6 km from the original point of tagging in the Black and Current rivers, Missouri, respectively. Measures of biomass and fish size indicated that adult shadow bass emigrated from the Current River to a large near-constant temperature spring (13.5◦ C) during cold winter months when river temperatures dropped below the spring temperatures. Individuals reentered the river during warm periods when river temperatures exceeded spring temperatures. During high use of the spring in cold periods, shadow bass in the spring had significantly higher relative stomach fullness and larger eggs than conspecifics in the river, suggesting that an energy subsidy was conferred on fishes that used the spring seasonally (Peterson and Rabeni 1996, reported as rock bass). Food: The shadow bass is primarily a benthic feeder. An extensive diet study in Missouri indicated that crayfish were by far the most important prey item in shadow bass >100 mm TL. Young-of-the-year initially relied on invertebrates, particularly chironomids and mayflies as prey, but began consuming crayfish at about 25 mm TL and increased consumption with growth. About 70% of usable energy of adult shadow bass was derived from consumption of crayfish. Shadow bass consumed crayfish species in proportion to their abundance in the river, were size selective for crayfish 30 to 44 mm in length, and showed no seasonal shifts in diet. Fish, primarily stonerollers, and other invertebrates, particularly mayflies and stoneflies, were additional, but less important, adult diet items (Probst et al . 1984; Rabeni 1992, reported as rock bass). A limited analysis of shadow bass diets in a small, sand-bottomed Gulf Coastal Plain stream in Louisiana indicated high consumption of benthic fish prey (e.g., darters, madtom catfish, shiners) and insects (e.g., dragonflies, stoneflies, caddisflies) but limited predation on crayfish (Viosca 1936). Diel activity and feeding studies are unavailable, but the absence of shadow bass at night from their daytime haunts suggests a nocturnal component in activity and perhaps foraging (or at least a nocturnal shift in habitat use) (Probst et al . 1984). Reproduction: Maturity is reached at age 1+ and a minimum size of 87 mm TL in females and 108 mm TL in males (C. S. Schieble, University of New Orleans, personal communication). Nest building has not been described, but an extensive examination of reproductive biology is available for southern populations in Lake Pontchartrain, Pearl River, and Mississippi River tributaries (C. S. Schieble, University of New Orleans, personal communication). Based on ovarian condition and ovary to body weight ratios, southern populations have a protracted spawning period extending from January or February to May or June, corresponding to water temperatures ranging from 15 to 26◦ C. Peak ovarian condition occurs at about 23◦ C. Mature ovarian eggs average 0.98 mm diameter (range, 0.56–1.7 mm), suggesting a somewhat smaller average mature ova size than in rock bass, but maximum sizes are comparable (Gross and Nowell 1980). Two size classes of vitellogenic ova are reported in mature females, and these are present from January through May, suggesting production of multiple batches of eggs. At a mean size of about 120 mm SL, a female can potentially produce 1311 mature eggs (range: 161 eggs at 85 mm SL to 4113 eggs at 156 mm SL) in a single spawning event. Peak female ovary to body weight ratios average 4.1% in February and March and 2.7% in March through May. Female ovary to body weight ratios, mean total ova, and mean ova diameters decrease substantially in June and subsequent months (C. S. Schieble, University of New Orleans, personal communication). Nest associates: None known. Freshwater mussel host: None documented, but see account on A. constellatus. Conservation status: The shadow bass appears to be secure throughout its range (Warren et al . 2000), but is considered vulnerable in Louisiana (NatureServe 2006) where it is confined to the southeastern portion of the state. Increased sedimentation and turbidity in formerly clear, relatively fast-flowing Gulf Coastal Plain and Mississippi Alluvial Valley streams could and likely have reduced available habitat for this species (Pflieger 1997; C. S. Schieble, University of New Orleans, personal communication).
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Similar species: Color pattern of sides of adult Ozark bass and rock bass (>100 mm TL) are irregularly arranged freckles or rows of blackish spots, lacking the usually conspicuous, alternating light and dark blotches of adult shadow bass. Juveniles of all three species are similarly patterned (Pflieger 1997). Systematic notes: Patterns of differentiation in the Ozark populations of A. ariommus and its sister species, A. rupestris, can render identification difficult, irrespective of whether morphological criteria or allozyme-derived genetic data are used. Some suggest that the patterns of differentiation indicate a north-to-south cline between A. rupestris and Ozarkian A. ariommus populations that are indicative of conspecificity, but the observed patterns are confounded by known or suspected introductions of both species into various drainages in the region. For example, populations of Ambloplites in the Gasconade River and Charette Creek (both Missouri River drainage) display allozyme-derived genetic distances intermediate between A. rupestris and A. ariommus, which are likely attributable to past introductions (Koppelman et al . 2000). Even in naturally occurring populations, intermediacy is not positive proof of conspecificity of A. rupestris and A. ariommus because long-term evolutionary retention of ancestral polymorphisms after divergence of sister species is common in centrarchids (Near et al . 2005). Further, morphological differences between the two species in the Ozarks are supported (e.g., cheek and breast scales, adult color patterns) (Koppelman et al . 2000). At this time, field identification of A. ariommus in the Ozarks appears to be best accomplished on the basis of adult body coloration, body depth to length ratio, aspects of squamation, and geography (Pflieger 1997; Koppelman et al . 2000). Notwithstanding the Ozarkian populations, extensive morphological comparisons and limited population sampling of allozymes indicate that A. ariommus is polytypic. Populations in drainages of the Florida Panhandle and perhaps the Mobile Basin may be distinct (Cashner 1974; Koppelman et al . 2000), but resolution of the nature of the differentiation awaits a rangewide phylogeographic analysis of the species. Importance to humans: The shadow bass has many desirable qualities as a sport fish although the relatively small maximum size limits angler interest in some parts of its range. The species readily takes a lure or natural baits and is a popular catch for anglers using ultralight gear or fly rods in streams and rivers of the Coastal Plain of Mississippi and the Ozark and Ouachita Mountains of Missouri and Arkansas (Robison and Buchanan 1984; Probst et al . 1984; Ross 2001). Creel surveys in the Pascagoula and Pearl rivers of Mississippi indicated that shadow bass constituted 1% and 0.6% of the total catch by weight, respectively (Ross 2001). The flavor and texture of the flesh of the shadow bass is similar to other centrarchids such as spotted bass and bluegill (Viosca 1936).
13.4.2 Ambloplites cavifrons Cope 13.4.2.1 Roanoke bass Characteristics: See generic account for general characteristics. Relatively large, elongate body; body depth >0.41 of SL. Eye large, diameter about 0.25 of head length. Body pattern similar to that of A. rupestris but with freckled pattern (scattered, dark brown spots) on side of body and head. Adults with unique color pattern of numerous iridescent gold to white spots on upper body and head. Preopercle strongly serrate at the angle. Dorsal fin elements, (22)23(24); anal fin elements, (16)17(18). Cheeks naked or incompletely scaled with small, deeply imbedded scales. Lateral line scales, (39)42 to 46(49); scale rows above lateral line, (8)9 or 10(12); scale rows below lateral line, (13)14 or 15(16); diagonal scale rows, 23 to 26(27); and breast scale rows, (26)30 to 34(36). One or two oval patches of teeth on tongue (Bailey 1938; Cashner 1974; Cashner and Jenkins 1982; Page and Burr 1991; Mabee 1993). Size and age: Typically reach 42 to 89 mm TL at age 1. Large individuals measure 250 to 296 mm TL, weigh 770 g, and reach age 4+ to 9+ (355 mm TL) (Smith 1971; Carlander 1977; Petrimoulx 1983; Jenkins and Burkhead 1994). World angling record, 620 g, Virginia (IGFA 2006). State records in Virginia and North Carolina are 1.12 and 1.13 kg, respectively. The Roanoke bass is the largest species in the genus with many plausible historical accounts of individuals weighing >1.0 kg (Jenkins and Burkhead 1994). Coloration: Numerous iridescent gold to white spots on upper side of body and head. Ground colors variable, ranging from olive to tan to black to cream or blends of lighter and darker shades. Lateral pattern may consist of parallel rows of black spots, formed by scales darkened at bases, producing a lined pattern or indistinct dark and light blotches. Sides transition to white to bronze on breast and belly. All fins with some degree of yellow pigment, but median fins tend to be
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more olive and may be mottled or barred. Membranes of anal fin of breeding males dusky to dark but lack dark marginal band (Cashner 1974; Cashner and Jenkins 1982; Page and Burr 1991). Sexual dimorphism in color is minimal, but during nest guarding and spawning, the male darkens intensively and the pale spots become more evident (Petrimoulx 1984). Native range: The Roanoke bass is endemic to the Neuse, Tar, Roanoke, and Chowan river drainages, North Carolina, and Virginia (Page and Burr 1991). Habitat: The Roanoke bass occurs across a broad range of stream types in the upper Coastal Plain, Piedmont, Blue Ridge, and Ridge and Valley. The species is most common in flowing, rocky, and sandy creeks and small to medium rivers above the Fall Line, where it is often associated with deep runs. Roanoke bass appear to frequent faster currents than congeners (Smith 1971; Petrimoulx 1983; Jenkins and Burkhead 1994). Food: The Roanoke bass is primarily a benthic feeder. Crayfish are the most important prey item for adults (>150 mm TL), augmented by small fish (e.g., darters, catfish, shiners) and various aquatic insects, particularly mayflies and caddisflies (Smith 1969, 1971; McBride et al . 1982; Petrimoulx 1983). Fish are less important in the diet in spring than in summer or fall, but overall, 75% of the food volume of adults consists of crayfishes, and the remaining 25% is primarily fishes (Petrimoulx 1983). Young fish (9 and maximal alkalinities >2000 mg/l in alklai lakes of Nebraska. Other centrarchids introduced in these habitats survived from a few hours to less than a month (McCarraher and Gregory 1970; McCarraher 1971). The species can reproduce in ponds with maximal pH and dissolved solids of 8.8 and 19,248 mg/l, respectively (Imler et al . 1975), and chloride–sulfate alkalinities of 17 ppt (McCarraher and Gregory 1970). Food: The Sacramento perch is a sluggish, slow-stalking, highly opportunistic suction-feeding carnivore (Vinyard 1982; Moyle 2002). It feeds primarily by “inhaling” organisms off the bottom or aquatic plants and by capturing zooplankton, fish, or emerging insects in midwater (Moyle et al . 1974). The species has numerous, long gill rakers that likely play an important functional role in the extended (90 mm TL) in an introduced population (Pyramid Lake, Nevada) switched almost exclusively to piscivory, but in many populations, microcrustaceans and aquatic insect larvae and pupae continue as important components of the adult diet (Moyle et al . 1974; Imler et al . 1975; Aceituno and Vanicek 1976). Reproduction: Maturity is reached at age 2 to 3+ at a minimum size of about 120 mm fork length (FL). Spawning occurs at water temperatures of 18 to 29◦ C and can extend from March through early August with peaks in late May to early June (Murphy 1948; Mathews 1962; McCarraher and Gregory 1970; Aceituno and Vanicek 1976; Moyle 2002). Published accounts of reproductive behaviors are few, somewhat inconsistent, and based on limited observations. Although some observations suggested definite male territory defense (about 40 cm diameter) without preparation of the substrate, more recent extensive observations indicate male digging of nests with the caudal fin and subsequent defense of obvious cleared, depressions (C. M. Woodley, University of California-Davis, personal communication). Territories and nests are often associated with vegetation or filamentous algae beds in shallow water (20–50 cm deep) and over substrates of mud, clay, or rocks; rock piles or other cover may also attract spawning individuals (Murphy 1948; Mathews 1962, 1965; Aceituno and Vanicek 1976; Moyle 2002; C. M. Woodley, University of California-Davis, personal communication). Nest preparation may span several days (Moyle 2002). Some observed nests were arranged linearly along shorelines, but others were suggestive of colonies (Murphy 1948; Aceituno and Vanicek 1976; Moyle 2002). Tail quivering occurs in territorial males, a behavior which appears distinct from the nest sweeping behavior of other centrarchids (caudal sweeping, Miller 1963; Mathews 1965). The male remains stationary over the nest with the head down and pectoral fins out and rapidly oscillates the tail back and forth in small arcs, at 3 to 5 oscillations per second, ending with the head up and nearly perpendicular to the nest. After several seconds the male rests, then repeats the behavior, which intensifies during courtship and spawning. Territorial males repeatedly repulse approaching females (Mathews 1965). After repeated attempts to repulse
Centrarchid identification and natural history
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the female (≤1 hour), the male swims stiffly to the ready female and nips at the vent (Moyle 2002). Pairs of Sacramento perch spend up to 30 minutes on the nest before spawning, during which time the male nips or nudges the female and both substrate bite, undulate, and contort their bodies, and jaw gape. Females may mate with more than one nesting male (Moyle 2002). In a natural setting, a male and female in the nest oriented broadside during spawning, but in opposite directions, unlike the head-to-head spawning position typical of other centrarchids. They made tight circles during gamete release as is typical of many centrarchids, but both the male and female tilted away from one another at the moment of release, another apparent departure from typical centrarchid gamete release (Mathews 1965; see also Bolnick and Miller 2006). Eggs are demersal, slightly adhesive, and upon deposition, adhere to surrounding vegetation or substrate in the bottom of the nest. Sacramento perch have among the smallest mature eggs among centrarchids (0.67 mm diameter) (Mathews 1962) and one of the highest batch fecundities among centrarchids (see Centrarchus macropterus and Pomoxis). Descriptive accounts indicate a unimodal distribution of mature or ripening ova sizes in mature females (Mathews 1962), suggesting release of a single batch of eggs. The relationship between number of mature eggs (Y) and TL (X) is described by the power function Y = 0.0279X2.6148 (n = 32, R2 = 0.89, data from Mathews 1962, FL converted to TL, see Aceituno and Vanicek 1976). At a mean size of 200 mm TL, a female can produce 29,003 mature eggs (range: 9820 eggs at 117 mm TL to 121,570 eggs at 330 mm TL, Mathews 1962). Hatching occurs in 51 hours and larval swim-up between 4 and 6 days at 22◦ C (Mathews 1962). From a single nest observation, male parental care is oft-cited as lasting only 3.5 days at water temperatures between 22 and 24◦ C, which is a short period of parental care relative to other centrarchids (Mathews 1965). More extensive observations at cooler water temperatures indicate that males stay at the nest for 5 to 7 days, apparently abandoning the nest only after larvae swim-up and move out of the nest area (Mathews 1962, 1965; C. M. Woodley, University of California-Davis, personal communication). Nest associates: None known. Freshwater mussel host: None known. Conservation status: Although tolerant of a range of physicochemical conditions, the distribution and abundance of native populations of the Sacramento perch has declined gradually since the nineteenth century. Declines are attributed to habitat alteration, embryo predation, and interspecific competition, particularly from nonnative centrarchids, such as bluegill and black crappie (Murphy 1948; Aceituno and Nicola 1976; Vanicek 1980; Marchetti 1999; Moyle 2002). In experiments with limited food resources, growth was depressed and habitat use shifted in the Sacramento perch in the presence of the more aggressive, dominating bluegill (Marchetti 1999). Native populations in the Pajaro and Salinas rivers and Clear Lake (Lake County) are extirpated (Gobalet 1990; Moyle 2002; Schwartz and May 2004). Within their native range the species persists primarily in ponds, reservoirs, and recreational lakes into which they were introduced, often upstream of native habitat (Moyle 2002). The species is considered of special concern in California rather than endangered because a few introduced populations appear secure (e.g., Garrison Reservior, Utah; Crowley Reservoir, California). However, even in many introduction sites in California and elsewhere, the species is uncommon, extremely rare, or extirpated (Moyle 2002; P. Crain and C. M. Woodley, University of California-Davis, personal communications; see section on native range). Similar species: The anal fin base of the white crappie and black crappie is about as long as the dorsal fin base, and the dorsal fin in these species has six to eight spines. Systematic notes: Archoplites interruptus is sister to the genus Ambloplites, and the Archoplites–Ambloplites pair are sister to Pomoxis (Roe et al . 2002; Near et al . 2004, 2005). Fossil representatives of the genus Archoplites are widespread west of the continental divide in Miocene to Early Pleistocene deposits (e.g., Idaho, Washington, Oregon, Utah, Nevada, and California) (Miller and Smith 1967; Smith and Miller 1985; Minckley et al . 1986; McPhail and Lindsey 1986; Near et al . 2005). Two other species, both extinct, are congeners: A. clarki Smith and Miller, from Miocene lacustrine deposits in northern Idaho (Smith and Miller 1985) and A. taylori Miller and Smith, from Late Pliocene to Early Pleistocene lacustrine deposits in southwestern Idaho (Miller and Smith 1967; Smith and Patterson 1994). Meristic variation among populations of A. interruptus is low, but some differences in color pattern exist (Hopkirk 1973; Moyle 2002). The population in Clear Lake probably is genetically distinct because of long isolation from other populations (Moyle 2002). Importance to humans: Historically, the Sacramento perch was one of the most common fishes caught by native peoples of California. In the late nineteenth century, 18,144 to 195,954 kg (40,000 to 432,000 lb) were sold annually in San Francisco (Gobalet and Jones 1995; Moyle 2002).
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Centrarchid fishes
13.6 Centrarchus macropterus (Lac´ep`ede) 13.6.0.2 Flier Characteristics: Deep, extremely compressed body, depth about half of SL. Small, supraterminal, oblique mouth, lower jaw projecting, supramaxilla moderate (2.1 to ≤3 times into length of maxilla), upper jaw not reaching past middle of eye. Eye large, diameter equal or greater than snout length. Large black teardrop. Interrupted rows of dark spots along the side. Juveniles (≤65 mm SL) with red-orange halo encircling black spot on posterior of soft dorsal fin. Opercle lacks flat extensions; opercular spot black. Preopercle posterior margin finely serrate. Long dorsal fin, 11 to 14 spines, 12 to 15 rays, 25 to 27 total; and long anal fin, 7 to 9 spines, 13 to 17 rays, 22 to 24 total. Dorsal fin base about 1.1 to 1.3 times longer than anal fin base. Spiny and soft dorsal fins continuous and smoothly rounded. Emarginate caudal fin. Long, pointed pectoral fin. Long, slender gill rakers, 30 to 40. Ctenoid scales. Lateral line scales 36 to 44; cheek scale rows, 4 to 7; branchiostegal rays, 7; pectoral rays, (12)13(14); vertebrae, 31(13 + 18). Teeth on entopterygoid, ectopterygoid, palatine (villiform), and glossohyal (tongue, two patches) bones (Bailey 1938; Page and Burr 1991; Mabee 1993; Jenkins and Burkhead 1994; Boschung and Mayden 2004). Size and age: Typically reach 55 to 72 mm TL at age 1. Large individuals measure 210 mm TL, weigh 156 to 197 g, and reach age 7+ to 8+ (maximum 250–356 mm TL) (Conley 1966; Geaghan 1978; Etnier and Starnes 1993; Jenkins and Burkhead 1994; Pflieger 1997). World angling record, 560 g, Georgia and North Carolina (IGFA 2006). Females can reach larger sizes and live longer than males (Conley 1966; Geaghan and Huish 1981). Coloration: Olive green to olive brown above; sides brassy yellow or silver with green and bronze flecks; rows of brown spots on sides forming horizontal lines. Brown-black spots on medial fins often form wavy bands or bars. Iris with vertical black bar continuing as tear drop. Young with four to five broad dark bars on side (Page and Burr 1991; Jenkins and Burkhead 1994; Pflieger 1997; Boschung and Mayden 2004). Native range: The flier occurs primarily on the Coastal Plain from the Potomac River drainage, Maryland, to central Florida, and west to the Trinity River, Texas. The species penetrates the Mississippi Embayment to southern Illinois and southern Indiana, where it occurs above the Fall Line (Page and Burr 1991). Habitat: The flier is a decidedly lowland species, inhabiting swamps, vegetated lakes, ponds, sloughs, and backwaters and pools of small creeks and small rivers. The species is usually associated with densely vegetated, clear waters (Page and Burr 1991; Jenkins and Burkhead 1994; Pflieger 1997; Boschung and Mayden 2004). Relative abundances were highest in hypoxic habitats in the Atchafalaya River Basin, Louisiana, where most fishes occurred in low relative abundances (Rutherford et al . 2001). The species also occurs in acid waters (pH 3.7 to 4.8), although growth appears to be diminished at low pH (Geaghan 1978); it is the most common sunfish in the acidic Okefenokee Swamp (Laerm and Freeman 1986). Movements of 12.7 km are documented, but ≥75% of individuals recaptured within 90 days of marking were found 1.2 of snout length. Six bold, black bars on sides, the first passes through the eye, the third extends dorsally through anterior spiny dorsal fin and ventrally through medial portion of pelvic fin, and the sixth through the caudal peduncle (often faint). Opercular spot dark with pale medial crescent. Rounded or slightly truncate caudal fin in young and juvenile, becoming truncate or slightly emarginate in adults. Long dorsal fin, (8)10(11) spines, 11 to 12 rays, usually 21 total, and short anal fin, 3 spines, (11)12 to 13(14) rays, 14 to 16 total. Dorsal fin continuous with deep notch between spines and rays. Dorsal fin base about 1.5 times longer than anal fin base. Dorsal and caudal fins not enlarged in breeding male. Pectoral fin narrow, somewhat pointed. Lateral line complete. Lateral scales, (23)25 to 29(32); cheek scale rows, (2)3(4); caudal peduncle scale rows, (16)18 to 21(22); pectoral rays, (9)11(13). Teeth present or absent on palatine bone (Bailey 1938; Page and Burr 1991; Mabee 1993; Jenkins and Burkhead 1994). Size and age: Typically reach 13 to 40 mm TL at age 1. Large individuals measure 40 to 60 mm TL (maximum 80 mm TL) and reach age 4+ (Schwartz 1961; Page and Burr 1991; Jenkins and Burkhead 1994). Length–weight relationships between males and females are similar in some populations (Schwartz 1961), but in a Delaware population females lived longer (age 3+) and reached larger maximum sizes (70 mm SL) than males (age 1+, 25 mm SL. Rounded caudal fin. Long dorsal fin, (7)9(11) spines, (10)11(13) rays, usually 21 total, and short anal fin, 3 spines, (10)10 to 11(12), 13 to 14 total. Dorsal fin continuous. Dorsal fin base about 1.5 to 1.7 times longer than anal fin base. Breeding male with enlarged second dorsal and anal fins and longest pelvic rays distally filamentous; female lacks enlarged fins and filamentous extensions. Pectoral fin rounded. Lateral line usually interrupted or incomplete. Lateral scales, (27)30 to 32(35); cheek scale rows, (3)4(5); caudal peduncle scale rows, (17)19 to 22(24); pectoral rays, (10)11 to 12(13). Teeth (cardiform) present on palatine bone (Bailey 1938; Peterson and Ross 1987; Page and Burr 1991; Mabee 1993; Jenkins and Burkhead 1994). Size and age: Reached 20 to 30 mm TL at age 1 in a Connecticut reservoir (Cohen 1977); age 0+ fish were 34 to 35 mm SL in October and 51 mm SL the following April in the Okefenokee Swamp (Freeman and Freeman 1985). Large individuals measure 55 mm TL (maximum 95 mm TL) and reach age 6+ (Cohen 1977; Page and Burr 1991). Males tend to live longer and grow slightly faster than females (Cohen 1977). Coloration: Dusky olive above, light below, with olive-black or five to eight black bars on the sides that may vary in distinctiveness. Rows of purple-gold crescentic flecks along side. Opercular spot black, bordered with iridescent goldgreen margin. Median fins dark with rows of blue to white spots. Breeding male, and to a lesser degree, breeding female with gold-green or blue flecks on head, body, and median fins, fin spines glowing white. Iris orange-red (Page and Burr 1991; Jenkins and Burkhead 1994). Aspects of subtle differences in coloration between E. obesus and E. gloriosus are summarized by Jenkins and Burkhead (1994). Native range: The banded sunfish occurs primarily on the Coastal Plain of Atlantic and Gulf Slope drainages from southern New Hampshire south of central Florida and west of the Perdido River drainage of Alabama (Page and Burr 1991; Boschung and Mayden 2004). Across the range, the species can be rare to relatively common (Smith 1985; Laerm and Freeman 1986; Jenkins and Burkhead 1994; Boschung and Mayden 2004; Marcy et al . 2005). An introduced population is established in the Black River drainage of Mississippi (Peterson and Ross 1987). Habitat: The banded sunfish inhabits heavily vegetated lakes, ponds, and sluggish sand- or mud-bottomed pools and backwaters of creeks and small to large rivers (Page and Burr 1991). The species is perhaps one of the most acid-tolerant fishes known (Gonzalez and Dunson 1987) and occurs in waters with pH 3.7 (e.g., New Jersey, Graham and Hastings 1984; Graham 1989; Georgia, Freeman and Freeman 1985). In multivariate studies in New Jersey, the banded sunfish was associated more strongly with acidic (pH 6.6–4.1), dystrophic habitats than either congener in lakes (Graham and Hastings 1984; Graham 1993) and in streams occurred most frequently between pH 5.0 and 4.5 (Zampella and Bunnell 1998). Individuals survived 2-week laboratory exposures to pH 3.5, and 60% of test individuals survived 3-week exposures to pH 3.3 after a gradual lowering from 3.5 over a 1-week period (Gonzalez and Dunson 1987). Growth was unaffected down to a pH of 3.75 (Gonzalez and Dunson 1989c). These findings suggest that the banded sunfish may have distinct
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competitive advantages over congeners and other sunfishes in low pH habitats (Gonzalez and Dunson 1991). Its tolerance of low pH is the result of complex adaptations for compensating for losses in body Na that would kill other fishes and involves the ability to limit branchial electrolyte permeability during acidic exposure (Gonzalez and Dunson 1987, 1989a,b,c). The gills of banded sunfish have a high affinity for Ca that reduces leaching by H+ and prevents high Na losses down to pH 3.5. In addition to limiting Na efflux, the species apparently can shift internal Na from osmotically inactive sources (e.g., bone) to plasma, which maintains Na concentrations of extracellular fluid. Although chronic acid exposure causes a large drop in body Na concentration (up to 52%, lethal to most fishes), these adaptations allow the banded sunfish to survive (Gonzalez and Dunson 1987, 1989a,b,c, 1991). Food: The banded sunfish, like its sister species the bluespotted sunfish, is an opportunistic forager on benthic, vegetational, and planktonic prey; adult diets are dominated by prey associated with submerged aquatic vegetation (Chable 1947; Cohen 1977; Graham 1989). Although diets overlap substantially between the two species, the banded sunfish gleans more vegetational prey and eats less benthic and planktonic prey than the bluespotted sunfish where the two co-occur (Graham 1989). Dominant adult food items are chironomid larvae (and other aquatic insects) and small crustaceans (cladocerans, copepods, amphipods). The young transition from a diet predominated by cladocerans, copepods, and chironomid larvae to the broader adult diet (Graham 1989). In late summer, young-of-the-year stomachs were nearly empty at dawn, but stomach fullness and digestion of prey indicated that individuals began feeding at dawn, paused between late morning and midday, and then fed continuously until dark (Graham 1986). Reproduction: Maturity is reached at age 2+ in females at a size of about 35 to 40 mm TL, but some smaller, age 1+ females are capable of spawning (Cohen 1977). Information on minimum size and age of maturity of males is lacking, but males are reproductively active by at least 59 mm TL (Harrington 1956). Gonadal development and associated nesting and spawning behaviors are controlled by increasing photoperiod and temperature (Harrington 1956). When males and females collected from ponds in fall were exposed in the laboratory to 15 hours of daylight and 21.7◦ C water temperature, ovary volume, ova size, testis volume, and male breeding colors developed rapidly (about 38 days), and nest building and spawning occurred. In contrast, in a parallel set of experiments at 21.7◦ C conducted under a fall photoperiod (9.2–11.6 hours daylight), individuals did not show gonadal enlargement or other reproduction-associated changes. In natural environments, spawning can be protracted. Gravid females and nuptial males occur from April to July in Virginia (Jenkins and Burkhead 1994), and capture of small young in Delaware suggests a late spring-through-summer breeding season (Wang and Kernehan 1979). In contrast, young-of-the-year only appeared in early June collections in a year-long sampling effort in the Okefenokee Swamp, Georgia (Freeman and Freeman 1985). Peak spawning and egg development occurred in June and July in a Connecticut reservoir at surface water temperatures of 23 to 27◦ C. Most details of reproductive biology, spawning behavior, and aspects of parental care are undocumented. In aquaria, breeding males establish territories, engage in threat postures and chasing, excavate depressional nests with their mouths, and vigorously defend the nest, eggs, and free-swimming larvae (Harrington 1956; Breder and Rosen 1966; Cohen 1977; Rollo 1994). One large male (52 mm SL) bred on 10 different days (of 26 days observed) and participated in 107 spawning acts under laboratory conditions (Harrington 1956). The interval between spawning acts was from 0 to 4 days. Mean fecundity, presumably based on total ova, increases with age (and size) ranging from 802 eggs at age 1 to 1400 eggs at age 6 (Cohen 1977). Mature ova are 0.6 mm in diameter. Fertilized eggs are adhesive and colorless, eggs hatch in about 3 days at 21.7◦ C, and larvae become free swimming about 5 days after hatching (Harrington 1956; Rollo 1994). Nest associates: None known. Freshwater mussel host: None known. Conservation status: Although not in danger of imminent extinction because of occupation of broad latitudinal range across many independent drainage systems, the banded sunfish is considered vulnerable to critically imperiled in many states within its range (New Hampshire, Rhode Island, Connecticut, Virginia, Alabama, Pennsylvania, New York) (Warren et al . 2000; NatureServe 2006). Similar species: See account on bluespotted sunfish. Systematic notes: See account on E. gloriosus.
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Importance to humans: Like congeners, the banded sunfish is popular among enthusiasts interested in keeping and rearing native fishes (Rollo 1994; Schleser 1998). Although perhaps underappreciated, the ability of the species to tolerate waters of relatively high acidity should increase scientific interest in the species.
13.8 Lepomis Rafinesque The genus Lepomis is a monophyletic clade of 13 species and is sister to the genus Micropterus (Near et al . 2004, 2005). The natural range encompasses most of eastern North America east of the Rocky Mountains, reaching northward to the Great Lakes, St. Lawrence River, and Hudson Bay drainages of Canada and eastward and southward in the Mississippi River Basin, Atlantic Slope, and Gulf of Mexico drainages west to the Rio Grande. Breeding males of some Lepomis are among the most colorful of all North American native fishes, and the reproductive habits of several species are among the best-studied and most fascinating within the fish fauna. The literature is extensive and only a brief overview is presented here and in the individual accounts. Lepomis share many features common to centrarchid reproduction. Males establish territories, excavate nests, fan, and guard eggs and defend newly hatched larvae until the swim-up stage. In addition, many Lepomis develop brilliant breeding colors and possess highly complex reproductive behaviors that can involve motor, visual, and auditory signals, and several species have evolved alternative mating strategies. Territorial breeding males excavate the typical circular depressional nest of other centrarchids, but many distinctive behaviors and combinations of behaviors are documented, often being associated with nest defense, courtship, or both. The male is faced with defending a nesting territory using agonistic behaviors and successfully mating with a female using courtship behaviors, motivations that necessarily shift from moment to moment, particularly in colonial nesters, and often appear in conflict (Keenleyside 1967; Steele and Keenleyside 1971; Ballantyne and Colgan 1978a,b,c). Males over nests display to nearby or approaching males and females using combinations of nest hovering, dashes to the surface and back to the nest, nest sweeping with the caudal fin, fin spreading, mouth gapes, jaw snaps, lateral displays (males side-by-side with fins erect), breast displays, substrate biting, and opercular spreads. Males most frequently rush toward an interloper with a quick retreat to the nest (thrust, Miller 1963), but if the intruder does not retreat, males display or actually ram, push, bite, or jaw grasp the other male. Males may also engage in rim circling, in which males repeatedly and rapidly circle their nest (e.g., over 100 circles in 30 minutes) with fins displayed (Miller 1963; Hunter 1963; Huck and Gunning 1967; Boyer and Vogele 1971; Avila 1976; Colgan et al . 1979; Lukas and Orth 1993). The act likely makes the male more conspicuous to females (Miller 1963; Avila 1976) but also serves as a territorial advertisement to other males (Colgan et al . 1979). In courtship, as a spawning-ready Lepomis female approaches a male’s nest, the male performs courtship circles by darting from the nest with fins spread, encircling the female and leading her toward the nest (Keenleyside 1967; Boyer and Vogele 1971; Avila 1976; Ballantyne and Colgan 1978a,b,c; Gross 1982). The male may courtship circle many times in rapid succession until the female follows him to the nest or leaves (Miller 1963; Keenleyside 1967). Augmenting the motor behaviors and breeding colors developed on the body and head, males of some species also have exaggerated opercular flaps. The ear flaps (or ear tabs) are species specific in orientation, size, and color patterns and serve as sex ornaments (secondary sexual characteristics) that play a complex role in mate choice, species recognition, and aggression between rival males (Keenleyside 1971; Colgan and Gross 1977; Stacey and Chiszar 1977). Opercle flaring directed at females is frequent in courting males (Keenleyside 1967), and the flap apparently signals to the female the species, condition, and quality of the male (Childers 1967; Goddard and Mathis 2000). Females prefer males with larger opercular flaps (e.g., Lepomis megalotis), and larger flaps increase the probability of a male in attaining and holding central nesting sites in a colony, where females spawn preferentially relative to peripheral nests (e.g., Lepomis macrochirus) (Gross and MacMillan 1981; Cˆot´e and Gross 1993; Goddard and Mathis 1997; Ehlinger 1999). Aggressiveness and dominance also are closely linked to the opercular flap. Males of at least some Lepomis appear to assess the resource-holding power of rivals by their opercular flap size (Goddard and Mathis 2000). Out of age, size, and seven morphological features in male bluegill, opercular flap size was the only feature that corresponded significantly with male rank in a breeding territory dominance hierarchy in experimental tanks (Ehlinger 1999). Some territorial, breeding male Lepomis further augment motor and visual reproductive signals with sound. On sighting a female near his nest, a nesting male rushes toward her and back toward his nest while producing a series of gruntlike
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sounds (bluegill, green sunfish, longear sunfish, and redspotted sunfish) or popping sounds (pumpkinseed and redear sunfish) (Gerald 1971; Ballantyne and Colgan 1978a,b,c). The sounds are also produced as males attack other males intruding into their nesting territory or in noncourtship agonistic contexts (Ballantyne and Colgan 1978a,b,c). Sound production is attributed to manipulation of the pharyngeal jaw pads, but in agonistic or courtship contexts is not associated with feeding (Ballantyne and Colgan 1978a,b,c). Sound characteristics suggest species specificity (Gerald 1971), and conspecific and heterospecific sounds elicit auditory brainstem responses in Lepomis (Wysocki and Ladich 2003), but individual variation in sound characteristics is high (Ballantyne and Colgan 1978a,b,c). Females are more responsive to conspecific than heterospecific sounds, but males respond to both (Gerald 1971; Ballantyne and Colgan 1978a,b,c). Sound production may facilitate location of nesting males by females in conditions of low visibility (Gerald 1971; Steele and Keenleyside 1971), but the behavior also appears to be part of a ritualized sequence of behaviors (e.g., jaw snaps and courtship circles), signaling that the male is both highly aggressively and sexually aroused (Ballantyne and Colgan 1978a). Alternative male reproductive strategies are highly evolved in Lepomis (Gross 1982; Jennings and Philipp 1992a; Philipp and Gross 1994; Avise et al . 2002). In a nest takeover strategy, large guardian males permanently displace small guardian males, or in nesting colonies, neighboring guardian males may intrude temporarily in another male’s nest to steal fertilizations with a female (Keenleyside 1972; Avila 1976; Dominey 1981; Gross 1982; Dupuis and Keenleyside 1988; Jennings and Philipp 1992b,c; DeWoody et al . 1998). Nesting male Lepomis habituate to the appearance of males on neighboring nests and become less aggressive toward them (Colgan et al . 1979), so unmated neighbors can more easily intrude and steal fertilizations (Keenleyside 1972; Jennings and Philipp 1992b). These strategies, however, appear to occur in relatively low frequencies (96%) of the young in their nests. Nest takeovers were rare, but 44% of assayed nests contained low percentages of offspring from nonguardian males, even though no sneaker male morphs were detected (DeWoody et al . 1998; DeWoody and Avise 2001). Intrusion by an ostensible female between a spawning pair (Lukas and Orth 1993) also suggests the possibility of sneaker males in some populations. Mature ovarian eggs range from 0.90 to 1.64 mm (mean 1.20 mm) (Sandow et al . 1975). The relationship between total number of mature ova (Y) and total length (X) is described by the linear function log Y = −3.8786 + 3.1628 log X (n = 79, R2 = 0.71, equation from Sandow et al . 1975). At a median size of 153 mm TL, a female can potentially produce 1074 mature eggs in a single batch (range: 435 at 115 mm TL to 6104 eggs at 265 mm TL). The adhesive, yellow to amber, fertilized eggs hatch in 3 days at 20 to 24◦ C. Newly hatched larvae are 4.6 to 5.1 mm TL, and most larvae are free swimming at 7.6 to 8.2 mm TL (Hardy 1978; Buynak and Mohr 1978; Yeager 1981). The guardian male vigorously defends the nest, eggs, and larvae from nest predators, may reduce foraging activity, and may cannibalize offspring in his own nest (Thorp et al . 1989; Lukas and Orth 1993; DeWoody et al . 2001). Nest associates: Dusky shiner, Notropis cummingsae (Fletcher 1993); swallowtail shiner, Notropis procne (Buynak and Mohr 1978); golden shiner, Notemigonus crysoleucas (Shao 1997). Freshwater mussel host: Putative host to Lampsilis teres, L. recta, and V. constricta (unpublished sources in OSUDM 2006). Conservation status: The redbreast sunfish is widespread and often abundant within its native range. It is considered vulnerable in Rhode Island, Massachusetts, and New York (Smith 1985; NatureServe 2006). In Massachusetts, it appears to have declined since the mid-1800s owing to changes in water quality or behavioral interactions with introduced species, especially the bluegill (Hartel et al . 2002). Similar species: Adult longear, northern longear, and dollar sunfishes have a shorter ear flap that is bordered by a white or orange edge, possess blue marbling or spots on the side of the adult, and lack distinct rows of red-brown spots on the upper side (Page and Burr 1991). Systematic notes: Lepomis auritus is sister to a clade inclusive of L. marginatus, L. megalotis, and L. peltastes (Near et al . 2004, 2005). Comparative studies of variation across the range of L. auritus are lacking. Importance to humans: The redbreast sunfish is a popular, sought-after sport fish in streams and rivers across most of the Atlantic Slope and eastern Gulf Coast (e.g., Suwannee River). On light tackle, redbreast sunfish offer excellent sport, being somewhat more aggressive, more surface oriented, and more active in cool waters than bluegill. The quality of the flesh is excellent and rated higher than that of Micropterus by some (Etnier and Starnes 1993; Jenkins and Burkhead 1994).
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13.8.2 Lepomis cyanellus Rafinesque 13.8.2.1 Green sunfish Characteristics: See generic account for general characteristics. Body deep, compressed, but elongate and thick relative to other Lepomis, depth 0.37 to 0.45 of SL. Mouth large, terminal, slightly oblique, supramaxilla small (>3 and ≤4 times length of maxilla), upper jaw extends well beyond anterior edge of eye, and in large individuals may extend to posterior edge of eye or beyond. Adult with dark spot at posterior base of soft dorsal and sometimes anal fin. Green to blue wavy lines on sides of snout, cheek, and opercle. Opercular flap stiff, short, black in center, edged in pale or yellow tinge that extends forward to form light borders above and below. Pectoral fin short and rounded, tip usually not reaching eye when laid forward across cheek. Long slender gill rakers, 11 to 14, longest about six times greatest width, thicker in large adults. Lateral line complete. Scales small. Lateral scales, (41)45 to 50(53); rows above lateral line, 8 to 10; rows below lateral line, 16 to 19; cheek scale rows, 6 to 9; caudal peduncle scale rows, 23 to 25; pectoral rays, 13 to 15. Pharyngeal arches narrow, strong, with small, thin, sharply pointed to conically blunt teeth. Teeth on palatine bone. No teeth on endopterygoid, ectopterygoid, or glossohyal (tongue, rarely a few teeth present) bones (Bailey 1938; Childers 1967; Trautman 1981; Becker 1983; Page and Burr 1991; Etnier and Starnes 1993; Mabee 1993). Size and age: Size at age 1 is highly variable among habitats and across latitudes, ranging from 30 to 165 mm TL (median 51 mm). Large individuals measure 150 to 225 mm TL, weigh 85 to 200 g, and attain age 5+ to 6+ (maximum 310 mm TL, age 10+) (Carlander 1977; Page and Burr 1991; Pflieger 1997; Quist and Guy 2001). World angling record, 0.96 kg, Missouri (IGFA 2006). Growth in mid-western prairie streams, where the species is common, is associated positively with abundance of instream wood, likely reflecting cover or food resources associated with wood (Quist and Guy 2001). Males may grow faster and perhaps live longer than females, but differences can be slight, becoming most apparent in individuals >100 mm TL (Hubbs and Cooper 1935; Carlander 1977). Coloration: Black, relatively short, ear flap with conspicuous light border. Wavy, often narrow, blue lines radiate from mouth across sides of snout onto cheek and opercle (often broken on opercle). Yellow, orange, or whitish margins on second dorsal fin, caudal fin lobes, anal fin, and pelvic fins, more prominent in breeding males. Blue-green above and on sides; iridescent, narrow, pale blue stripes on body scales interspersed with yellow metallic flecking; the blue stripes often broken into irregular mottling or spotting, especially posteriorly; sometimes with dusky bars on side. White to yellow belly (Hunter 1963; Page and Burr 1991; Etnier and Starnes 1993; Jenkins and Burkhead 1994). Native range: The green sunfish is native to the east-central United States, west of the Appalachians from the Great Lakes, Hudson Bay, and Mississippi River Basins from New York and Ontario to Minnesota and South Dakota and south to the Gulf Slope drainages from the Escambia River, Florida, and Mobile Basin, Georgia and Alabama, west to the lower Rio Grande basin, Texas, and northern Mexico (Page and Burr 1991; Miller 2005). The species has been widely introduced and is established over much of the United States including Atlantic and Pacific Slope drainages and Hawaii (Page and Burr 1991; Fuller et al . 1999). Introduced populations of green sunfish in Atlantic Slope and in western US waters are implicated in suppression and decline of native game and nongame fishes as well as frogs and salamanders (Lemly 1985; Fuller et al . 1999; Dudley and Matter 2000; Moyle 2002). Habitat: The green sunfish is a highly successful, aggressive, competitive species occurring in a variety of habitats including clear to turbid headwaters, sluggish pools of large streams, isolated, dry season–stream pools, and shallow shorelines of lakes, ponds, and reservoirs (Werner and Hall 1977; Werner et al . 1977; Capone and Kushlan 1991; Page and Burr 1991; Etnier and Starnes 1993; Taylor and Warren 2001; Smiley et al . 2005). In pond experiments, the presence of green sunfish induced dramatic shifts in foraging habitat and prey types in co-occurring congeners (Werner and Hall 1977, 1979). Green sunfish also invoke strong antipredator behaviors in aquatic insects and amphibians (e.g., Sih et al . 1992; Krupa and Sih 1998). The species is among the most tolerant of Lepomis to adverse conditions of high turbidity (400 m), and long-distance movements (>16 km) are
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documented (Funk 1957; Hasler and Wisby 1958; Kudrna 1965; Smithson and Johnston 1999). The green sunfish is also an adept disperser and “pioneer” species, rapidly colonizing streams recovering from seasonal drying or drought, moving into and out of seasonally inundated floodplain habitats, and often invading ponds or small lakes (Ross and Baker 1983; Matthews 1987; Kwak 1988; Capone and Kushlan 1991; Etnier and Starnes 1993; Taylor and Warren 2001; Moyle 2002; Adams and Warren 2005). Food: The adult green sunfish is a solitary ambush predator whose large mouth allows it to feed on larger food items at a given body size than most congeners (Sadzikowski and Wallace 1976; Werner and Hall 1977). The size-adjusted gape area of the species is the second largest within the genus (see L. gulosus; Collar et al . 2005a,b). The adult diet consists primarily of aquatic insects, particularly large odonate, mayfly, and beetle larvae; fish; crayfish; and terrestrial invertebrates, but a variety of other taxa are consumed (e.g., snails, and unusually, a bat) (Minckley 1963; Applegate et al . 1967; Etnier 1971; Sadzikowski and Wallace 1976; Werner 1977; Carlander 1977; Lemly 1985). Young green sunfish transition from an initial diet of microcrustaceans to larger invertebrates and at 50 to 99 mm TL increase consumption of crayfishes and fishes (Applegate et al . 1967; Mittelbach and Persson 1998). High volumes of plant material in stomachs are indicative of considerable foraging for invertebrates, such as odonate larvae, associated with vegetation (Etnier 1971; Sadzikowski and Wallace 1976). In laboratory studies, activity levels are largely diurnal, peaking at dusk and dawn, but the presence in stomachs of prey only available after dark indicates a nocturnal or at least crepuscular component to feeding (Etnier 1971; Beitinger et al . 1975; Langley et al . 1993). Green sunfish produce a chemical alarm substance that induces antipredatory behaviors in conspecifics, regardless of size. In contrast, chemical alarm cues from sympatric heterospecific fishes induce antipredator responses in juvenile green sunfish and foraging responses in adults (Golub and Brown 2003). Reproduction: Maturity is reached at age 1+ to 3+ at a minimum size of about 45 to 76 mm TL (Carlander 1977). The combined effects of increased photoperiod (15 hours) and rising temperature in spring control prespawning gonadal development (Kaya and Hasler 1972; Smith 1975). Under controlled photoperiods, temperature, and food availability, 6-month old individuals (60–100 mm TL) can be induced to spawn (Smith 1975). Spawning is protracted (mid-May to early August), with the initiation of spawning depending in part on latitude (Hunter 1963; Kaya and Hasler 1972; Carlander 1977; Pflieger 1997). Nest building and spawning begin as water temperatures increase to 20◦ C, and peak spawning occurs between about 20 and 28◦ C (Hunter 1963). Nesting activity decreases and gonadal regression occurs as water temperatures remain over 28◦ C for extended periods (Hunter 1963; Kaya 1973). Males excavate nests by caudal sweeping. Nests are about 31 cm in diameter and usually placed over gravel in open, shallow areas (4–35 cm water depth, rarely 100 cm). Within a population, small males nest later in the season and in shallower water than large males (Hunter 1963), and at similar latitudes, individuals from stunted populations become ripe 2 to 4 weeks later than nonstunted populations (Childers 1967). Nests may be widely spaced (up to 30 m apart) when population densities are low but can also be placed rim-to-rim in crowded colonies (Hunter 1963; Childers 1967; Pflieger 1997). Colony formation closely parallels that of other colonial-nesting Lepomis (e.g., Bietz 1981; Neff et al . 2004), but whether colonial nesting occurs in the absence of habitat limitation is not completely clear (Hunter 1963; Childers 1967; Pflieger 1997). Spawning events are synchronous in colonies, occurring at intervals of 8 to 9 days over the spawning season; males may nest five or more times in succession during this period, and females presumably participate in multiple spawning events (three to six) over the season (Hunter 1963). Nest-guarding males produce gruntlike sounds as part of courtship (Gerald 1971); other reported courtship, spawning, and nest defense behaviors appear typical for the genus (Hunter 1963; Childers 1967). During nest building and spawning, males are territorial, aggressive, and even combative toward other males, females, and nest predators; only the most persistent spawning-ready females are allowed into the nest. Activity of spawning males is intensified. For example, in a 10-minute period a guardian male completed five spawning acts, made ten defensive forays outside the nest, threatened his neighbor once, and rim-circled 39 times (Hunter 1963). During a given spawning event, females attempt to mate (and likely do mate) with multiple males, but appear most attracted to males that are already spawning. Occasional intrusions by an ostensible female between a spawning pair (Hunter 1963) suggest the presence of sneaker males in at least some populations, but alternative mating systems in green sunfish are unconfirmed. Mature ovarian eggs are 0.8 to 1.0 mm in diameter, and fertilized eggs are 1.0 to 1.4 mm in diameter (mean 1.23 mm) (Meyer 1970; Kaya and Hasler 1972; Taubert 1977). Depending on their size, females may carry 2000 to 10,000 eggs (Beckman 1952 in Moyle 2002), but little else is apparently known about fecundity. The adhesive, fertilized eggs hatch in 2.1 days at 23.8◦ C (1.3 days at 27.1◦ C) (Childers 1967). Newly hatched larvae are 3.6 to 3.7 mm TL, and, depending on temperature, larvae are free swimming for about 3 to 6 days after fertilization at 4.6 to 6.3 mm TL (Childers 1967; Meyer 1970; Taubert
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1977). Successful males guard and vigorously defend the nest, eggs, and larvae for 5 to 7 days, but earlier abandonment of nests is common (Hunter 1963). Nest associates: Red shiner, Cyprinella lutrensis (Pflieger 1997); redfin shiner, Lythrurus umbratilis (Hunter and Wisby 1961; Hunter and Hasler 1965; Snelson and Pflieger 1975; Trautman 1981; Johnston 1994a,b; Pflieger 1997); golden shiner, N. crysoleucas (suspected, Pflieger 1997); Topeka shiner, Notropis topeka (Pflieger 1997). Freshwater mussel host: Confirmed host to A. ligamentina, Anodonta suborbiculata, Elliptio complanata, Glebula rotundata, Lampsilis altilis, Lampsilis bracteata, Lampsilis cardium, Lampsilis higginsii , Lampsilis hydiana, L. reeveiana, Lasmigona complanata, Ligumia subrostrata, L. recta, Megalonaias nervosa, P. grandis, V. iris, Villosa vibex , and U. imbecillis (Young 1911; Lefevre and Curtis 1912; Tucker 1927, 1928; Stern and Felder 1978; Trdan and Hoeh 1982; Parker et al . 1984; Waller and Holland-Bartels 1988; Howells 1997; Barnhart and Roberts 1997; Haag et al . 1999; O’Dee and Watters 2000). Putative host to A. plicata, Lampsilis radiata, Lasmigona compressa, S. undulatus, Toxolasma lividus, and Toxolasma parvus, (unpublished sources in OSUDM 2006). Conservation status: Although abundant in few natural habitats (e.g., Pflieger 1997; Quist and Guy 2001), the green sunfish is widespread and stable within its native range. Similar species: Other Lepomis lack yellow-orange edges on the fins and the black spot at posterior base of the dorsal fin (except the bluegill) and have a smaller mouth (except the warmouth). The bluegill has long, pointed pectoral fins, and the warmouth has dark red-brown lines radiating posteriorly from the eye, mottling on the side, and a small patch of teeth on the tongue (Page and Burr 1991). Systematic notes: Lepomis cyanellus forms a sister pair with L. symmetricus, and the pair represents the second largest and the smallest Lepomis, respectively (Near et al . 2004, 2005). Comparative studies of variation across the range of L. cyanellus are lacking. Importance to humans: The green sunfish rarely reaches a size of interest to anglers other than children. Because of its propensity to invade, overpopulate, stunt, and compete with other fishes in ponds or small lakes, green sunfish are considered a pest by those attempting to maintain quality bluegill-bass sport fisheries. The species is commonly used by anglers as live bait on trotlines, set hooks, and jugs for catfishes. Hybrids between a female green sunfish and a male bluegill (known as “hybrid bream”) are cultured and stocked in ponds to create put-and-take fisheries. The hybrids are aggressive, fast growing, and easy to catch, and if properly managed, produce excellent results (Ross 2001).
13.8.3 Lepomis gibbosus (Linnaeus) 13.8.3.1 Pumpkinseed Characteristics: See generic account for general characteristics. Body, deep, compressed, often almost disk-like, depth about 0.40 to 0.53 of SL. Mouth moderate, terminal, slightly oblique, supramaxilla absent, upper jaw extends almost to, or to, anterior edge of eye. Wavy blue lines on cheek and opercle of adult. Bold dark brown wavy lines or orange spots on soft dorsal, anal, and caudal fins. Opercular flap stiff, short, with black center bordered in white or yellow with a prominent red (males) to yellowish (females) semicircular spot at posterior edge (often pale or yellowish in young). Pectoral fin long, sharply pointed, usually reaching far past eye when laid forward across cheek. Short, thick gill rakers, about 12; scarcely longer than wide. Lateral line complete. Lateral scales, (35)37 to 44(47); rows above lateral line, 6 to 8; rows below lateral line, 12 to 15; cheek scale rows, 3 to 6; caudal peduncle scale rows, 17 to 21; pectoral rays, 11 to 14. Pharyngeal arches extremely broad, heavy with large rounded, molariform teeth. Teeth present or absent on palatine bone. No teeth on endopterygoid, ectopterygoid, or glossohyal (tongue) bones (Scott and Crossman 1973; Trautman 1981; Becker 1983; Page and Burr 1991; Mabee 1993; Jenkins and Burkhead 1994). Size and age: Size at age 1 is highly variable among habitats and across latitudes, ranging from 15 to 99 mm TL (median 40 mm). Large individuals measure 150 to 225 mm TL, weigh about 150 to 200 g, and attain age 6 to 9+ (maximum 400 mm TL, age 10+) (Carlander 1977; Page and Burr 1991; Fox 1994). World angling record, 0.63 kg, New Mexico (IGFA
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2006). Pumpkinseed populations sympatric with bluegill show increased early growth rates, despite reduced resources, relative to populations allopatric with bluegill, providing evidence for counter-gradient evolutionary selection for rapid growth (Arendt and Wilson 1997, 1999). Older males tend to be larger than same-age females, and subtle differences in body form occur between male and female pumpkinseed (Deacon and Keast 1987; Brinsmead and Fox 2002). Coloration: Ear flap black with light border, marked with bright red or yellow-orange spot on posterior edge. Wavy, usually wide, blue lines radiate from mouth across sides of snout onto cheek and opercle of adult. Many bold dark brown wavy lines or orange spots on second dorsal, caudal, and anal fins. Olive above and on sides with many gold and yellow flecks. Adults blue-green, spotted with orange; dusky chainlike bars mark sides of young and adult female; white to red-orange below (Page and Burr 1991). Native range: The pumpkinseed is native to Atlantic Slope drainages from New Brunswick south to the Edisto River, South Carolina, and to the Great Lakes, Hudson Bay, and upper Mississippi River Basins from Quebec and New York west to southeast Manitoba and North Dakota and south to northern Kentucky and Missouri. The species has been widely introduced and is established over much of the United States and southern Canada, including some Pacific Slope drainages (Scott and Crossman 1973; Page and Burr 1991; Fuller et al . 1999; Moyle 2002). Habitat: The pumpkinseed inhabits vegetated lakes and ponds and quiet vegetated pools of creeks and small rivers (Page and Burr 1991). Lake- and stream-dwelling populations differ in subtle aspects of body morphology (e.g., pectoral fin length), differences attributed to adaptation to lentic versus lotic environments (Brinsmead and Fox 2002). Juvenile and adult pumpkinseed tend toward lengthy occupancy of home activity areas (about 11 m2 to 1.12 hectares, respectively) and can home to those areas when displaced (Shoemaker 1952; Hasler et al . 1958; Kudrna 1965; Reed 1971; Fish and Savitz 1983; Wilson et al . 1993; Coleman and Wilson 1996; McCairns and Fox 2004). Food: The pumpkinseed is a highly specialized molluscivore, feeding primarily on snails by crushing them between heavy pharyngeal jaw bones that are equipped with molariform teeth, enlarged muscles, and specialized neuromuscular adaptations (Lauder 1983a,b, 1986; Hambright and Hall 1992; Wainwright and Lauder 1992; Huckins 1997). Adults also feed heavily on dipteran, mayfly, and caddisfly larvae and beetles, and also ingest cladocerans, amphipods, isopods, ostracods, larval odonates, and terrestrial invertebrates (Seaburg and Moyle 1964; Sadzikowski and Wallace 1976; Keast 1978; Laughlin and Werner 1980; Deacon and Keast 1987; Huckins 1997; Jastrebski and Robinson 2004). Young age-0 fish (>18 mm TL) consume a diet predominated in biomass by zooplankton and chironomids (Hanson and Qadri 1984), and at least in pond experiments, their combined predatory effects can change zooplankton composition (Hambright and Hall 1992). As they grow from 35 to 100 mm TL, the young transition gradually from a diet of soft-bodied littoral invertebrates to high numbers of snails (Keast 1978; Mittelbach 1984a; Keast and Fox 1990; Osenberg et al . 1992; Huckins 1997). Full development of the pharyngeal snail-crushing apparatus of pumpkinseeds depends on repeated, consistent consumption of snails (Bailey 1938). Pharyngeal bones and musculature associated with snail crushing are substantially reduced in individuals in snail-poor lakes relative to individuals from snail-rich lakes (Wainwright et al . 1991; Mittelbach et al . 1992; Osenberg et al . 2004). In the summer, peaks in feeding occur in late afternoon and at dawn with reduced but notable feeding after midnight (Keast and Welsh 1968). In the fall, daylight feeding is low and feeding peaks occur between 2000 and 0400 hours (Johnson and Dropkin 1993). In summer, age-0 pumpkinseed feed from shortly after sunrise until sunset (Hanson and Qadri 1984). Periodic infrared videography of foraging pumpkinseed over 8 months revealed frequent nocturnal foraging, mediated by a switch from benthic picking during daylight to zooplanktivory at night (Collins and Hinch 1993). In support of these field observations, laboratory experiments indicate volumes searched and feeding rates on zooplankton decrease at light intensities ≤10 lux (Hartleb and Haney 1998). Pumpkinseeds produce a chemical alarm substance that induces antipredatory behaviors in conspecific juveniles (95 mm SL) (Marcus and Brown 2003; Golub et al . 2005). Response of juveniles to alarm cues was diminished under weakly acidic conditions (pH 6.0) (LeDuc et al . 2003). Pumpkinseed also respond to chemical alarm cues of largemouth bass (and ostariophysan alarm chemicals), but the response is mediated by size and habitat complexity. Under conditions of low to intermediate habitat complexity, large pumpkinseed (>80 mm SL) exhibit foraging responses and small pumpkinseed antipredator responses to bass chemical alarm cues. In highly complex habitat, both large and small pumpkinseed show antipredator responses to bass chemical alarm cues (Golub et al . 2005).
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Lake-dwelling pumpkinseeds show subtle intra- or interpopulation differences in body form (e.g., body depth, fin length, gill raker spacing) that are strongly associated with specializations for pelagic or littoral feeding (Robinson et al . 1996; Robinson and Schluter 2000; Brinsmead and Fox 2002; Gillespie and Fox 2003; Jastrebski and Robinson 2004; McCairns and Fox 2004). Intermediate forms occur in both habitats but show reduced fitness in growth and body condition (Robinson et al . 1996). Evidence from parasite analyses and strong site fidelity in pelagic and littoral zone pumpkinseed morphs suggest that trophic divergence and habitat segregation come into play early in the life history and could potentially affect gene flow (Robinson et al . 2000; Jastrebski and Robinson 2004; McCairns and Fox 2004). Intrapopulation morphological divergence between trophic morphs occurs across a relatively broad geographic region (Robinson et al . 2000; Gillespie and Fox 2003; Jastrebski and Robinson 2004). Divergence is expressed in the absence of open-water competitors (i.e. bluegill or other Lepomis) (Robinson et al . 1993), but may also be mediated by complex interactions of a number of ecological factors (Robinson et al . 2000). Reproduction: Maturity is reached at age 1+ to 4+ at 65 to 130 mm TL. Within a population, females may mature earlier and at smaller sizes than males (Carlander 1977; Fox and Keast 1991; Fox 1994; Danylchuk and Fox 1994; Fox et al . 1997). Age and size at maturity, onset and duration of spawning, size of eggs, and energy allocated for reproduction are plastic, varying in different, but proximate habitats (e.g., beaver ponds and nearby lakes, adjacent lakes) or regionally. Trade-offs among somatic growth and reproductive timing and allocation are linked to energy limitations, resource uncertainty in highly variable environments, and presence of other Lepomis (Deacon and Keast 1987; Fox and Keast 1991; Danylchuk and Fox 1994; Fox 1994; Fox et al . 1997). Spawning is protracted (early May to August), the initiation of spawning depending in part on latitude and population size structure (Burns 1976; Carlander 1977; Danylchuk and Fox 1994; Fox and Crivelli 1998). Gonadal development in both sexes accelerates as water temperatures warm to 12.0◦ C and photoperiod lengthens to 13.5 hours (Burns 1976). A combination of long photoperiod (16 hours) and warm temperature (25◦ C) induces nest-building behaviors in males (Smith 1970). Nest building and spawning begin as water temperatures increase to 17◦ C, and peak spawning occurs between about 20 and 22◦ C, but continues to at least 26◦ C (Miller 1963; Fox and Crivelli 1998; Cooke et al . 2006). Onset of spawning is later and the spawning season is longer in stunted than in nonstunted populations (Danylchuk and Fox 1994). Males excavate nests by caudal sweeping and uprooting and carrying away plants; conspecific or other centrarchid nests are often appropriated or reused (Ingram and Odum 1941; Miller 1963). Nests are 30 to 80 cm in diameter, at water depths of 18 to 50 cm (rarely >1 m), and often near simple cover (e.g., log, stump, boulder). Sand or small rocky substrates are chosen most often for nest sites, but a variety of substrates are used (Breder 1936; Ingram and Odum 1941; Colgan and Ealey 1973; Popiel et al . 1996). Nests are usually solitary (>1 m apart), but groups of two or three nests may be rim to rim (Ingram and Odum 1941; Miller 1963; Clark and Keenleyside 1967; Colgan and Ealey 1973). Nest-guarding males produce popping sounds as part of courtship of females and aggression toward conspecific males and other Lepomis (Gerald 1971; Ballantyne and Colgan 1978a,b,c). Other reported courtship, spawning, and nest defense behaviors appear typical for the genus (e.g., aggressive displays, courtship circles, rim circling) (Miller 1963; Steele and Keenleyside 1971; Colgan and Gross 1977; Colgan et al . 1981; Becker 1983; Clarke et al . 1984). Sneaker males are documented for pumpkinseed (Gross 1979), but in one surveyed population, guardian males sired about 85% of the larvae in their nests (range, 43–100%) (Rios-Cardenas and Webster 2005). Mature ovarian eggs average 1.11 mm diameter (Gross and Sargent 1985), but 0.6 to 1.0 mm and 0.8 to 1.2 mm diameters are ranges reported for fertilized or fertilized and water-hardened eggs, respectively (Hardy 1978; Cooke et al . 2006). Female batch fecundity increases with weight, but varies significantly among populations (Deacon and Keast 1987). The relationship between batch fecundity (Y) and total weight (X) is described by the linear function, log10 Y = −0.0592 + 1.9461 log10 X (n = 37, R 2 = 0.20, one of four equations from Deacon and Keast 1987). At 48 g (128 mm TL), a female can potentially produce 5455 mature eggs in a single batch (range: 2451 at 20 g and 98 mm TL to 10,633 eggs at 126 g and 184 mm SL, respectively). The white to transparent, adhesive, fertilized eggs hatch in about 3 days at 18 to 22◦ C, larvae at hatching are 2.6 to 3.1 mm TL, and larvae reach swim-up at about 5.2 mm TL, some 4 days after hatching (Miller 1963; Colgan and Gross 1977; Hardy 1978). The cycle for the successful guardian male typically takes 10 days (range 6–15 days) with 2 days for territory establishment and nest construction, three for spawning and egg guarding, four for larval guarding, and one for fry dispersal and nest abandonment. Territoriality and aggressiveness in guardian males is highest during egg guarding and early larval stages, diminishing as larvae grow (Colgan and Gross 1977; Colgan and Brown 1988; Cooke et al . 2006). Males may lose on average 6.3% of their body weight from spawning to fry dispersal (Rios-Cardenas and Webster 2005). Females can participate in one to six spawning periods (average two to three) over a 7- to 8-week period, during which an estimated
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12 to 40% of prespawning body mass is allocated to reproduction (Fox and Crivelli 1998). In lakes, fry apparently initially disperse offshore but return to littoral habitats in late summer (Keast 1978; Brown and Colgan 1984, 1985a; Mittelbach 1984a; Rettig 1998). Nest associates: Golden shiner, N. crysoleucas (Shao 1997). Freshwater mussel host: Confirmed host to Alasmidonta varicosa, P. grandis, and U. imbecillis (Trdan and Hoeh 1982; Fichtel and Smith 1995). Putative host to Alasmidonta undulata, A. plicata, E. complanata, L. radiata, Lampsilis siliquoidea, L. reeviana, Lasmigona costata, L. recta, P. cataracta, and S. undulatus (unpublished sources in OSUDM 2006). Conservation status: The pumpkinseed is secure across most of its native range but is considered critically imperiled in Manitoba and vulnerable in Illinois (NatureServe 2006), which include the northwestern and southern peripheries of its native distribution, respectively (Page and Burr 1991). Similar species: All other Lepomis have shorter, rounded pectoral fins, except the redear sunfish and bluegill. The redear sunfish and bluegill lack bold spots on the second dorsal fin and wavy blue lines on the gill cover (Page and Burr 1991). Systematic notes: Lepomis gibbosus is basal to a clade consisting of L. microlophus, and the sister pair L. punctatus– L. miniatus (Near et al . 2004, 2005). Based on shared behavioral and morphological specializations for snail crushing, L. gibbosus was proposed previously as sister to L. microlophus (Bailey 1938; Mabee 1993). Frequencies of nuclearencoded allozyme loci across populations in four east-central Ontario watersheds revealed low genetic variability, but populations were significantly substructured genetically. The patterns in genetic variation are congruent with hypothesized post-Pleistocene recolonization routes (Fox et al . 1997). Comparative studies of variation across the entire range of L. gibbosus are lacking, but anal and dorsal ray counts and differences in size and age at maturity show east to west differences (Scott and Crossman 1973; Fox et al . 1997). Importance to humans: Although not often reaching a size of interest to many anglers, the pumpkinseed can contribute substantially to the sport fishery catch in northern lakes (e.g., Minnesota, Eddy and Underhill 1974; Wisconsin, Becker 1983), at least historically contributed to the Great Lakes commercial fishery catch (Scott and Crossman 1973), and is an easy and delightful catch for young anglers. The flesh is white, flaky, sweet, and delicious, comparable to that of the bluegill. The species can be taken in late afternoons with light tackle on live bait, small dry flies, poppers, or wet fly trout patterns (Scott and Crossman 1973; Eddy and Underhill 1974; Becker 1983). The pumpkinseed is important ecologically, forming part of the food for many predatory fishes including important game fishes (e.g., black basses, walleye, yellow perch, and muskellunge) (Scott and Crossman 1973). Among northern North American freshwater fishes, the pumpkinseed is among the most striking in beauty and color (Jordan and Evermann 1923; Becker 1983). Because of their color and ease of keeping and breeding, the species is a prized aquarium fish in Europe (Goldstein 2000).
13.8.4 Lepomis gulosus (Cuvier) 13.8.4.1 Warmouth Characteristics: See generic account for general characteristics. Body relatively thick, robust, somewhat elongate, depth 0.4 to 0.5 of SL. Large, terminal oblique mouth, lower jaw projecting slightly, supramaxilla moderately large (>2 to ≤3 times length of maxilla), upper jaw extending well beyond anterior edge of eye to center of eye or beyond in adults. Dark red-brown lines (3–5) radiating posteriorly from snout and red eye. Opercular flap short, stiff, black with paler and often red-tinged border. Pectoral fin short and rounded, tip usually not reaching eye when laid forward across cheek. Long, thin gill rakers, 9 to 13, longest about four (adults) to six (young) times the greatest width. Lateral line complete. Lateral scales, 36 to 48; rows above lateral line, 6 to 9; rows below lateral line, 12 to 15; cheek scale rows, 5 to 7; caudal peduncle scale rows, 19 to 23; pectoral rays, 12 to 14. Pharyngeal arches narrow with bluntly conical teeth. Teeth on endopterygoid, ectopterygoid, palatine (villiform), and glossohyal (tongue, one patch) bones (Bailey 1938; Birdsong and Yerger 1967; Trautman 1981; Becker 1983; Etnier and Starnes 1993; Mabee 1993; Jenkins and Burkhead 1994; Boschung and Mayden 2004).
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Size and age: Size at age 1 is highly variable among habitats and across latitudes, ranging from 25 to 155 mm TL (median 55.5 mm TL). Large individuals measure 150 to 200 mm TL, weigh about 200 g, and attain age 5 to 7+ (maximum 310 mm TL, age 8+) (Carlander 1977; Page and Burr 1991). World angling record, 1.1 kg, Florida (IGFA 2006). Coloration: Ear flap short, black with yellow edges and posterior red spot (adult). Dark red-brown lines radiating from back of red eye. Olive brown above; dark brown mottling on back and upper side; often 6 to 11 chainlike dark brown bars on sides; cream to light yellow below; dark brown spots (absent on young) and wavy bands on fins. Breeding male boldly patterned on body and fins with a bright red-orange spot at base of second dorsal fin and black pelvic fins (Page and Burr 1991). Young and juveniles usually with a distinctive purplish sheen. Native range: The warmouth is native to the Great Lakes and Mississippi River Basin from western Pennsylvania to Minnesota and south to the Gulf of Mexico and the Atlantic and Gulf drainages from the Rappahannock River, Virginia, to, but apparently not including, the Rio Grande, Texas, New Mexico, and Mexico (Page and Burr 1991; Miller 2005). The species is an apparent recent (ca. 1966) natural immigrant in the waters of southern Ontario, where it is naturalized (Crossman et al . 1996). The warmouth has been introduced widely and is established over much of the United States, including some Pacific Slope drainages (Fuller et al . 1999; Moyle 2002). Habitat: The warmouth inhabits vegetated lakes, ponds, swamps, reservoirs, and quiet waters of slow-flowing streams, being most common, and often abundant, in lowland areas and rare in uplands (Larimore 1957; Holder 1970; Guillory 1978; Page and Burr 1991; Snodgrass and Meffe 1998). Individuals are most often solitary and usually associated with areas of dense vegetation, root wads, stumps, overhanging banks, or rock cavities over silt or mud substrates (Larimore 1957; Loftus and Kushlan 1987). Smaller warmouth (200 mm TL) often feed almost exclusively on crayfishes (Guillory 1978). Young warmouth transition from an initial diet of microcrustaceans to invertebrates (e.g., midge and caddisfly larvae) and at about 75 mm TL begin increasing use of the larger prey dominating the adult diet (Larimore 1957; Germann et al . 1974; Desselle et al . 1978; Guillory 1978). Dawn and dusk samples in the summer suggest that most feeding occurs at or before dawn with little feeding in the afternoon (Larimore 1957). Reproduction: Maturity is reached at ages 1+ to 2+ at 57 to 152 mm TL (Larimore 1957; Germann et al . 1974; Guillory 1978). Spawning is initiated as water temperatures approach 21◦ C (as low as 15◦ C) and is protracted (April or May to July or August) with female ovary to body weight ratios peaking in late May to early June as water temperatures reach 27 to 29◦ C (Larimore 1957; Germann et al . 1974; Guillory 1978). Males excavate nests in a few hours by caudal sweeping, and depending on the time spent by the male, the nest may be a rather shapeless oval depression (about 10 cm × 20 cm) with only loose silt swept away or a deep, symmetrical circular depression (45 cm diameter, 13 cm deep). Nests are constructed at water depths of 15 to 152 cm (most 4 m apart), but if habitat is limiting nests may be closely
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spaced (Carr 1940; Larimore 1957; Childers 1967). Courtship and spawning behaviors (based primarily on aquarium observations) appear typical for the genus (e.g., male aggressive displays, jaw gapes, opercular flares), but warmouth apparently do not rim circle; other than egg fanning by the male, no detailed observations are available on nest care or nest defense behaviors. During active courtship of a female, the body of a male becomes bright yellow and the eyes blood red in color, the change in colors requiring only 5 to 10 seconds. Only when the female is ready to lay eggs will she allow the male to guide her to the nest. In aquaria, a nest-guarding male will ultimately kill an unresponsive female (Larimore 1957). During paired circling of the nest (female near the center, male outside), the female jaw gapes a few times, violently jerks her body, and releases about 20 eggs while simultaneously thumping the male on the side in an apparent signal for him to release sperm. These behaviors are repeated sequentially for about 1 hour with brief pauses in between bouts, at which time males may use caudal sweeping to mix eggs into the substrate (Larimore 1957). Mature ovarian eggs (waterhardened) average 1.01 mm in diameter (Merriner 1971a). Mature females contain two or more egg class sizes throughout the spawning season (Larimore 1957; Germann et al . 1974). Batch fecundity increases with female size. The relationship between batch fecundity (Y) and total length (X) is described by the linear function, log10 Y = −1.6108 + 2.4859 log10 X (data from mean number of mature eggs of nine length classes, R2 = 0.85, Germann et al . 1974). At 195 mm TL, a female can potentially produce 12,078 mature eggs in a single batch (range: 6825 eggs at 155 mm TL to 20,238 eggs at 240 mm SL, respectively). Another estimate of batch fecundity is much lower (i.e. log10 Y = 0.1619 + 1.418 log10 X, where X is SL, Guillory 1978). The fertilized eggs are pale, amber-colored, and adhesive, hatching in about 1.5 days at 25.0 to 26.4◦ C (71.1 hours at 22.6◦ C, 33.9 hours at 26.9◦ C, and 32.5 hours at 27.3◦ C). Larvae at hatching are 2.3 to 2.9 mm TL and reach swim-up at about 4.7 to 7.6 mm TL, some 3 to 5 days after hatching (Larimore 1957; Childers 1967). After leaving the nest, young apparently do not form schools, but hide themselves in dense vegetation or other cover. Likewise, juvenile warmouth do not aggregate in large groups (Larimore 1957). Nest associates: Bluehead shiner, Pteronotropis hubbsi (Fletcher and Burr 1992). Freshwater mussel host: Confirmed host to A. suborbiculata, L. subrostrata, Toxolasma texasensis, and U. imbecillis (Stern and Felder 1978; Barnhart and Roberts 1997). Putative host to T. parvus (unpublished sources in OSUDM 2006). Conservation status: The warmouth is currently stable over most of its range (Warren et al . 2000; NatureServe 2006). Peripheral populations in Pennsylvania and West Virginia are considered imperiled, and recently naturalized populations in Ontario are listed as critically imperiled (NatureServe 2006), although the necessity for the latter status has been questioned (Crossman et al . 1996). Similar species: The green sunfish lacks dark lines radiating posteriorly from eye, lacks teeth on the tongue, and has a dark spot at the posterior base of the second dorsal fin (Page and Burr 1991). Systematic notes: Lepomis gulosus is basal to the sister pair L. symmetricus and L. cyanellus (Near et al . 2004, 2005). Mitochondrial DNA analyses revealed distinct eastern and western populations of L. gulosus, occurring along the Atlantic Slope through Florida to eastern tributaries of Mobile Basin and from the Tombigbee River westward, respectively (Bermingham and Avise 1986). L. gulosus has a checkered taxonomic and nomenclatural history (summary in Berra 2001), but comparative studies of variation across the range of the species are lacking. Importance to humans: Over much of its range, the warmouth is taken most often by bream or crappie anglers but usually not in abundance. Even so, warmouth can comprise a large part of the sport fish catch in habitats like the Okefenokee Swamp, Georgia, or Reelfoot Lake, Tennessee (Larimore 1957; Germann et al . 1974). Warmouth are quick to take an artificial lure or live bait. The species is an excellent table fish, the flavor and texture of the flesh being judged as intermediate between the bluegill and the largemouth bass (Larimore 1957).
13.8.5 Lepomis humilis (Girard) 13.8.5.1 Orangespotted sunfish Characteristics: See generic account for general characteristics. Body moderately deep, compressed, slab-sided, depth 0.38 to 0.45 of SL. Mouth moderately large, oblique, supramaxilla absent, upper jaw extends to, or just beyond, anterior edge of eye. Orange or red-brown wavy lines on cheek and opercle in adults. Opercular flap moderate to long (in adults),
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very flexible, usually angled upward with black center and wide, white to pale green, conspicuous border (flushed with orange in breeding males). Pectoral fin short and rounded, tip usually not reaching eye when laid forward across cheek. Moderately thin gill rakers, 10 to 15, longest about five times greatest width. Enlarged, elongate sensory pits on preopercle and head between eyes, pits larger than any other Lepomis, width of each pit about equal to distance between pits. Lateral line complete or incomplete. Lateral scales, 32 to 42; cheek scale rows, 5; pectoral rays, 13 to 15. Pharyngeal arches narrow with sharply pointed teeth. Teeth on palatine bone. No teeth on endopterygoid, ectopterygoid, or glossohyal (tongue) bones (Bailey 1938; Trautman 1981; Becker 1983; Mabee 1993; Ross 2001; Boschung and Mayden 2004). Size and age: Size at age 1 is highly variable among habitats and across latitudes, ranging from 23 to 86 mm TL (median 45 mm TL). Large individuals measure 75 to 125 mm TL, weigh 3 times and ≤4 times length of maxilla), upper jaw reaches posteriorly from beyond anterior of eye to just about center of eye. Wavy blue lines on cheek and opercle of adult. Opercular flap long, flexible (flared at end in large individuals), usually oriented horizontally (adult) or slanting upward (young), black in center with white edges, lower and upper edges of equal width, bordered above and below by blue line. Pectoral fin short and rounded, tip usually not reaching eye when laid forward across cheek. Short, thick, knoblike gill rakers, 12 to 14, longest about equal (adults) to twice (young) greatest width. Lateral line complete. Lateral scales, (31)36 to 48(50); rows above lateral line, (5)6 to 8(9); rows below lateral line, (11)14 to 15(19); cheek scale rows, (4)5 to 6(8); caudal peduncle scale rows, (16)18 to 23(25); pectoral rays, (11)13 to 14(15). Pharyngeal arches narrow with sharply pointed teeth. No teeth on endopterygoid, ectopterygoid, palatine, or glossohyal (tongue) bones (Bailey 1938; Barlow 1980; Trautman 1981; Mabee 1993; Boschung and Mayden 2004). Size and age: Size at age 1 is highly variable among habitats and across latitudes, ranging from 21 to 114 mm TL (median 47 mm TL). Individuals rarely exceed 155 mm TL or 100 g, and few live beyond age 6+ (maximum about 240 mm TL, 227 g, and age 9+) (Bacon 1968; Carlander 1977; Page and Burr 1991; Etnier and Starnes 1993; Jennings and Philipp 1992c). World angling record, 0.79 kg, New Mexico (IGFA 2006). Parental males grow faster than females (Carlander 1977; Jennings and Philipp 1992c). Coloration: Ear flap long, black in adult, edged in white, bordered above and below by blue lines. Numerous, wavy blue lines on sides of snout, cheek, and opercle. Young with olive back and side speckled with yellow flecks, often with
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chainlike bars on sides, white below. Adult dark red above, bright orange below, marbled and spotted with blue; clear to orange and blue, unspotted fins. Breeding males are among the most brilliantly colored North American fishes, with contrasting bright reddish orange and blue body, red eye, orange to red median fins, and blue-black pelvic fins (Page and Burr 1991). Nape with reddish stripe in upper Arkansas and Missouri River populations, and at least some populations in the upper White River, Missouri, lack the light border on the ear flap (Pflieger 1971; Barlow 1980; Goddard and Mathis 1997). Native range: The longear sunfish is native to the Mississippi River Basin west of the Appalachian Mountains from Indiana west to eastern Illinois and south to the Gulf of Mexico and to Gulf Slope drainages from the Choctawhatchee River, Florida, west to the Rio Grande, Texas, southern New Mexico, and northeastern Mexico (Page and Burr 1991; Miller 2005). The species is generally common, and often the most abundant Lepomis in upland or clear streams throughout its range. The species has expanded its range in recent decades north and westward in the Missouri River, Missouri, as a likely result of clear water conditions imposed on that system by upstream reservoirs (Pflieger 1997). The longear sunfish has been introduced sparingly outside its native range and is established in the upper Ohio River basin (New and Kanawha, above the Falls, rivers), the Atlantic Slope (Potomac River drainage and Maryland Coastal Plain), upper Rio Grande (New Mexico), and perhaps, the Pacific Slope of Mexico (Rio Yaqui) (Fuller et al . 1999; Miller 2005). Habitat: The longear sunfish inhabits rocky and sandy pools of headwaters, creeks, and small to medium rivers (Page and Burr 1991) and can thrive along shorelines of reservoirs (Bacon 1968; Gelwick and Matthews 1990; Bettoli et al . 1993; Etnier and Starnes 1993; Pflieger 1997). In some rivers, the longear sunfish can be the most abundant centrarchid (Gunning and Suttkus 1990). The species is tolerant of low DO (e.g., 100% survival at 34◦ C) (Matthews 1987; Smale and Rabeni 1995a,b; Beitinger et al . 2000). In streams, many individuals use restricted home activity areas (200 m) interhabitat and long-distance (140 mm TL. At 40 to 140 mm TL, small shoal bass transition from diets dominated by aquatic insect larvae (e.g., mayflies) to increased consumption of fish and crayfish (Wright 1967; Wheeler and Allen 2003). Reproduction: Females reach maturity at minimum sizes of 152 to 189 mm SL and age 2+, but most mature at age 3+ (Wright 1967; Hurst 1969; Hurst et al . 1975). On the basis of occurrence of ripe, partially spent, or recently spent females and observations in ponds, spawning occurs from April to May (perhaps into June) at water temperatures from 18.0 to 26.0◦ C. Ripe, presumably prespawning, females are taken at temperatures as low as 14.4◦ C in early April (Wright 1967; Hurst 1969; Smitherman and Ramsey 1972; Williams and Burgess 1999). Nests are circular depressions about 30 to 92 cm in diameter and 5 to 15 cm deep. In streams, nests are located in shallow water (20–45 cm deep) of pools upstream of riffles or in eddies adjacent to shoals, and in culture ponds, nests were excavated at water depths of 76 to 130 cm over clay, soft clay rubble, or plant roots (Wright 1967; Hurst 1969; Williams and Burgess 1999). Males reportedly vigorously guard the nest (Williams and Burgess 1999). Observations of a single spawning pair indicated an apparently typical Micropterus spawning sequence that lasted about 45 minutes and resulted in deposition of about 1000 large (2-mm diameter), amber-colored, adhesive eggs. While over the nest, the pair assumed a blotched coloration of dark green vertical bars on a background color of bronze. Other nests contained 500 to 3000 ova (Williams and Burgess 1999). Fecundity increases with female size but is not well quantified. The number of eggs (unclear whether total or mature) in five mature females ranged from 5396 eggs at 314 mm SL and 884 g to 21,799 eggs at 442 mm SL and 2314 g (Wright 1967). Eggs hatch in about 2 days at 21.1◦ C (Smitherman and Ramsey 1972), and yolk-sac larvae, averaging 4.4 mm TL, form tight aggregations in the nest bottom. The larvae reach swim-up about 7 days after hatching and disperse about 12 to 14 days after hatching (Smitherman and Ramsey 1972; Williams and Burgess 1999). Nest associates: None known. Freshwater mussel host: None known. Conservation status: The shoal bass is vulnerable throughout its native range (Warren et al . 2000). The species is considered critically imperiled in Florida, imperiled in Alabama, and vulnerable in Georgia (NatureServe 2006). In the Chattahoochee River, the shoal bass has disappeared from most of the main channel and declined in tributaries because of impoundments eliminating shoal habitats, increased sedimentation, and water quality degradation. Its former distributional
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extent in the Apalachicola and Flint rivers is also reduced by impoundments and channel dredging (Williams and Burgess 1999; Johnston 2004). Similar species: Superficially similar to redeye bass and spotted bass. Shoal bass (92% of specimens) lack a tooth patch on the tongue (versus oval to elongate patch in spotted bass and redeye bass). In adult shoal bass, the anterior half to two-thirds of the body has dark, vertically elongated, midlateral blotches that are separated by lighter areas approximately equal to the width of the blotch (versus irregular to more quadrate blotches in redeye bass); blotches usually confluent to form a midlateral stripe in spotted bass. Shoal bass also lack white outer edges on the caudal fin (present in redeye bass) and have higher caudal peduncle scale counts (Page and Burr 1991; Gilbert 1992a; Williams and Burgess 1999). Systematic notes: Micropterus cataractae is a member of a “Gulf of Mexico” clade of Micropterus, including all other Micropterus except M. dolomieu and Micropterus punctulatus (Kassler et al . 2002; Near et al . 2003, 2004). Relationships within the clade are not well resolved with M. cataractae placed as basal to the entire clade, sister to Micropterus coosae, sister to Micropterus notius, or basal to a clade inclusive of M. notius, M. p. henshalli,Micropterus treculi , and M. salmoides + Micropterus floridanus (Kassler et al . 2002; Near et al . 2003, 2004). Importance to humans: Shoal bass are the signature fish of a productive sport fishery in the Flint River, Georgia, particularly in the upper river (Davis 2006). Anglers wade fish the shoals using fly rods and crayfish-like flies or light to medium spinning gear with a variety of spinners, crayfish imitations, popping bugs, or other bass lures. The fast water habits of the shoal bass, a restricted native range, a scrappy fighting ability, and the propensity to take a fly and dive into the rocks, all combine for an exciting and specialty black bass catch. Supplemental stocking of shoal bass is being undertaken to augment the population in the lower Flint River (Davis 2006).
13.9.2 Micropterus coosae Hubbs and Bailey 13.9.2.1 Redeye bass Characteristics: See generic account for general characteristics. Elongate body, depth 0.20 to 0.24 of TL, increasing with size. Mouth large, terminal, lower jaw slightly projecting, upper jaw extends little or not at all beyond posterior edge of eye. Outline of spinous dorsal fin curved. Juncture of soft and spiny dorsal fins slightly emarginate, broadly connected. Shortest dorsal spine at emargination of fin, usually >0.75 times length of longest spine. Dorsal soft rays, usually 12, 11 to 14; anal soft rays, usually 10, 9 to 11. Gill rakers, (6)7 to 8. Lateral scales, (58)67 to 72(77); rows above lateral line, (7)8 to 9(13); rows below lateral line, (11)14 to 17(21); cheek scale rows, (8)12 to 13(16); caudal peduncle scale rows, (24)26 to 30(31); pectoral rays, (13)15 to 16(17). Small splintlike scales on interradial membranes at anal and second dorsal fin bases (>60 mm SL). Pyloric caeca, usually unbranched, 7 to 12. Teeth present or absent on glossohyal (tongue) bone (Hubbs and Bailey 1940; Ramsey and Smitherman 1972; Turner et al . 1991; Williams and Burgess 1999; Kassler et al . 2002). Size and age: Averages 49 to 63 mm TL (range, 38–68 mm) at age 1 in streams. Growth in ponds and reservoirs can be much higher (≥125 mm TL at age 1) (Parsons 1954; Gwinner et al . 1975; Catchings 1979; Barwick and Moore 1983). Young-of-the-year (22–25 mm TL) stocked in forage-supplemented ponds in June reached 134 mm TL by midDecember (Smitherman and Ramsey 1972; Smitherman 1975) and in some reservoirs individuals average 122 to 125 mm TL at age 1 (Barwick and Moore 1983). Few redeye bass reach 325 mm TL, exceed 225 g, and attain age 5+ to 7+ (maximum about 470 mm TL, 1.44 kg, and age 10+) (Parsons 1954; Smitherman 1975; Carlander 1977; Barwick and Moore 1983; Page and Burr 1991; Etnier and Starnes 1993; Boschung and Mayden 2004; OutdoorAlabama 2006). Redeye bass are perhaps the slowest growing Micropterus. The maximum size attained even in the fastest-growing reservoir populations suggests genetically based size limitations (Barwick and Moore 1983; Moyle 2002). Coloration: Uniquely, among all Micropterus, the outer margins of the caudal fin lobes in redeye bass are narrowly depigmented (in life iridescent white or frosted orange in color, may be less obvious in large individuals) (Ramsey 1975). Color above olive to deep bronze. Back to lateral midline marked with dark, vertically elongate, diamond-shaped to irregularly quadrate blotches, most evident in young, fading with age. Rows of dark spots usually evident on lower sides. Yellow-white ventral area. Iris characteristically red. Breeding males with aqua-blue to blue-green cast on lower half of head and ventral area. Young-of-the-year soft dorsal fin, caudal fin, and front of anal fin tinged brick red to orange; caudal
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fin lacks sharply contrasting tricolored pigmentation (Ramsey and Smitherman 1972; Page and Burr 1991; Turner et al . 1991; Etnier and Starnes 1993; Mettee et al . 1996; Boschung and Mayden 2004). Native range: The redeye bass is native above the Fall Line from the Savannah, Altamaha, and Chattahoochee rivers and the upper Mobile Basin (Coosa, Cahaba, Tallapoosa, and Black Warrior rivers) in North Carolina, South Carolina, Georgia, Tennessee, and Alabama (Page and Burr 1991; Williams and Burgess 1999). The native or introduced status of the species in the Santee River drainage, North and South Carolina, is uncertain (Warren et al . 2000), but preliminary genetic analyses suggest that the population(s) in the Saluda River is introduced (F. C. Rohde personal communication, Division of Marine Fishes, North Carolina). From about 1940 through the 1960s, the species was introduced outside its native range and is now established in tributaries of the Tennessee and Cumberland rivers, Tennessee and Kentucky, and in several drainages in California (Fuller et al . 1999; Moyle 2002). Although often debated as native rather than introduced (e.g., Clay 1975; Koppelman and Garrett 2002), established populations in Martins Fork Cumberland River, Kentucky, were introduced deliberately by state fisheries personnel around 1950 from stock obtained in Georgia (Burr and Warren 1986). In Tennessee and Cumberland river streams, introduced redeye bass have hybridized extensively and likely introgressed with native smallmouth bass (Turner et al . 1991; Pipas and Bulow 1998). Some superabundant stream populations of redeye bass developed after introductions in California, where the species is associated with declines of native minnows, suckers, salamanders, and ranid frogs (Fuller et al . 1999; Moyle 2002). Habitat: The redeye bass inhabits rocky, small upland creeks and small to medium upland rivers, where it is associated with pools, boulders, undercut banks, and water willow beds (Parsons 1954; Page and Burr 1991; Pipas and Bulow 1998; Moyle 2002). The species can be common even in the smallest headwater stream where few other fish and no other Micropterus occur (Parsons 1954; Ramsey 1975; Pipas and Bulow 1998). The redeye bass has been viewed traditionally as potentially providing a fishery in waters too cool and small for other Micropterus but too warm for trout (e.g., Parsons 1954; Carlander 1977). These conditions, however, are not prerequisites for establishment of thriving redeye bass populations in nonnative habitats (Pipas and Bulow 1998; Moyle 2002). Indirect evidence suggests that redeye bass make large upstream migrations to tributaries to spawn in the spring (and conversely downstream fall migrations to winter habitat) (Parsons 1954). Redeye bass are generally intolerant of ponds and most reservoirs (Parsons 1954; Wood et al . 1956; Webb and Reeves 1975; Moyle 2002; but see Barwick and Moore 1983). Food: The redeye bass is an opportunistic carnivore, feeding from the surface to the bottom. The summer diet in streams consists primarily of terrestrial insects and crayfish. To a lesser extent, stream-dwelling redeye bass also consume small fishes (e.g., minnows and darters), aquatic insects, and salamanders (Parsons 1954; Smitherman 1975; Gwinner et al . 1975). Large redeye bass (>216 mm TL) in oligotrophic reservoirs in South Carolina are primarily piscivorous (Barwick and Moore 1983). Reproduction: Maturity is reached at a minimum size of 120 mm TL at age 3+ in females and age 4+ in males in streams, but faster growing pond-cultured individuals matured at age 1+ (Parsons 1954; Smitherman 1975). Spawning extends from April to early July as water temperatures reach 18 to 21◦ C (Parsons 1954; Smitherman and Ramsey 1972; Gwinner et al . 1975). Practically nothing is published on male or female reproductive behaviors, and overall knowledge about the reproductive biology of redeye bass is at best sketchy. Nests are shallow, circular depressions in coarse gravel at the heads of pools (Parsons 1954). Fertilized, water-hardened eggs average 3.5 mm in diameter (Smitherman and Ramsey 1972). Relationships between female size and fecundity are unquantified. Two females of 145 and 205 mm TL contained 2084 and 2334 eggs, respectively (Parsons 1954). Eggs hatch in about 2 days at 22.8◦ C; yolk-sac larvae are 6.0 mm TL, and larvae are free swimming at 7 to 8 mm TL about 5 days after hatching (Smitherman and Ramsey 1972). An anecdotal account suggests that fry school for a short time relative to most Micropterus (Parsons 1954). In a culture pond, complete breakup of schools occurred at 16 to 25 mm TL about 14 days after swim-up, but school breakup began as early as 6 days after swim-up (Smitherman and Ramsey 1972). Nest associates: None known. Freshwater mussel host: Confirmed host to L. altilis, Lampsilis perovalis, V. nebulosa, and V. vibex (Haag and Warren 1997; Haag et al . 1999). Conservation status: The redeye bass is secure throughout its range (Warren et al . 2000), but native populations on the periphery of the range are considered vulnerable (Tennessee) or critically imperiled (North Carolina) (NatureServe
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2006). Obversely, the past introduction and establishment of redeye bass outside its native range now threatens the genetic integrity of populations of native Micropterus (Turner et al . 1991; Pipas and Bulow 1998). Similar species: See accounts on Suwannee bass and spotted bass. Differs from all other Micropterus in having the outer margins of the caudal fin lobes narrowly depigmented (iridescent white or frosted orange in life) (Ramsey 1975; Page and Burr 1991). Systematic notes: Micropterus coosae is a member of a “Gulf of Mexico” clade of Micropterus, including all other Micropterus except M. dolomieu and M. punctulatus (Near et al . 2003, 2004). Relationships within the clade are not well resolved with M. coosae placed as basal to the clade, sister to M. cataractae, sister to M. punctulatus henshalli (the Alabama spotted bass), or basal to M. notius, M. treculi, and M. salmoides + M. floridanus (Kassler et al . 2002; Near et al . 2003). Similarities in form, color, behavior, and ecology led most morphological taxonomists to relate M. coosae to M. dolomieu or M. punctulatus (e.g., Hubbs and Bailey 1940; Ramsey 1975). Data from nuclear-encoded allozyme loci and mitochondrial DNA reveal significant genetic substructuring among populations now known as redeye bass and strongly suggest the existence of multiple, and perhaps specifically distinct, evolutionary lineages (Kassler et al . 2002; Koppelman and Garrett 2002). The evolutionary relationships among populations of redeye bass, and of redeye bass to other Micropterus, particularly the Alabama spotted bass (see account on M. punctulatus), await thorough genetic evaluation. Importance to humans: The attractive redeye bass is regarded as a somewhat wary, but scrappy fighter in small, wadeable streams, where it provides an exciting catch on ultralight gear combined with small lures and spinners, popping bugs and flies, or natural bait (Parsons 1954; Etnier and Starnes 1993). In its small stream habitat, redeye bass populations can provide a minimal catch-and-release fishery, but slow growth rates limit establishment of harvestable stream fisheries (Pipas and Bulow 1998).
13.9.3 Micropterus dolomieu (Lac´ep`ede) 13.9.3.1 Smallmouth bass Characteristics: Elongate, slightly compressed body, depth 0.18 to 0.28 of TL, decreasing with size. Mouth large, terminal, lower jaw slightly projecting, upper jaw extends at least to below center of eye but not beyond posterior edge of eye. Outline of spinous dorsal fin curved. Juncture of soft and spiny dorsal fins slightly emarginate, broadly connected. Shortest dorsal spine at emargination of fin, usually >0.5 times the length of the longest spine. Dorsal soft rays, usually 13 or 14, 10 to 15; anal soft rays, usually 11, 9 to 12. Gill rakers, 6 to 8. Lateral scales, (64)69 to 77(81); rows above lateral line, (10)12 to 13(15); rows below lateral line, (16)19 to 23(32); cheek scale rows, (13)15 to 18(20); caudal peduncle scale rows, (26)29 to 31(33); pectoral rays, (13)16 to 17(18). Small splintlike scales on interradial membranes at anal and second dorsal fin bases (>60 mm SL). Pyloric caeca, unbranched, about 10 to 15. Teeth present or absent on glossohyal (tongue) bone (Bailey 1938; Hubbs and Bailey 1938, 1940; Smitherman and Ramsey 1972; Turner et al . 1991; Kassler et al . 2002). Size and age: Size at age 1 is highly variable among habitats and across latitudes and ranges from 40 to 188 mm TL (median 92 mm TL) (Beamesderfer and North 1995). Large individuals can exceed 400 mm TL, weigh 1.5 to 2.5 kg, and attain age 6+ to 12+ (maximum 686 mm TL, 5.2 kg, and age 14+) (Scott and Crossman 1973; Carlander 1977; Paragamian 1984; Page and Burr 1991; Weathers and Bain 1992; Beamesderfer and North 1995; MacMillan et al . 2002). World angling record, 4.93 kg, Tennessee (IGFA 2006). Growth rates are similar between males and females (Carlander 1977). Coloration: No dark lateral band. Dark brown with numerous bronze markings on scales, often with 8 to 16 indistinct vertical bars on a yellow-green to brown side. Olive brown with bronze specks above, yellow to white below. Iris usually reddish. Large male is green-brown to bronze with dark mottling on back and dark vertical bars on the side. Young (4 m in their activity areas, and at least some individuals occupy distinctive diurnal and nocturnal activity areas (Emery 1973; Savitz et al . 1993; Cole and Moring 1997). In Lake Opeongo, Ontario, smallmouth bass use the largest recorded summer home ranges among centrarchids. Average postnesting home range area is 247 ha for males and 409 ha females, but core use areas (50% use) are smaller (38.4 ha) and similar between sexes. Individual male summer home ranges show high coincidence from year to year, indicating that males in the lake return from nesting areas to the same home ranges over multiple years (Ridgway and Shuter 1996; Ridgway et al . 2002). Daytime movements within these large home ranges are extensive, averaging 4.8 km over 6- to 16-hour periods (about 483 m/h), but there is little activity at
Centrarchid identification and natural history
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night (Ridgway et al . 2002). The differences in home range size estimates among smallmouth bass in different lakes may be attributable to methods used to estimate home range (e.g., Savitz et al . 1993; Cole and Moring 1997; Ridgway et al . 2002) but may also reflect differences in resource availability (e.g., forage, cover) or in population-specific adaptations. Riverine smallmouth bass also show high persistence in relatively small areas throughout the summer months, but fall movement to winter habitats varies among populations (review by Lyons and Kanehl 2002). In a Missouri stream, postspawning home ranges and intrapool movement of adults were greater in summer (0.09 to 0.67 ha, up to 980 m/d at 27.5◦ C) than in winter (0.06 to 0.22 ha, 120 m/d at 4◦ C), but fish generally used the same stream sections in winter and summer, moving elsewhere only during the spawning season (Todd and Rabeni 1989). In small Ouachita Mountain streams, interpool movement of smallmouth bass in summer was high, with 35% of marked individuals moving among adjacent pools over a 3-day observation period (Lonzarich et al . 2000). Similarly, recolonization rates after complete removal were high; pool populations reached pre-removal abundances in 40 days (Lonzarich et al . 1998). Some populations of riverine smallmouth bass, particularly those in areas with severe winters, make fall migrations of several to over 100 km to wintering habitats (usually to downstream bodies of water) (e.g., Langhurst and Schoenike 1990; Peterson and Rabeni 1996; Cooke et al . 2000a; Lyons and Kanehl 2002; Schreer and Cooke 2002). Movement to wintering areas can involve numerous short movements with rest periods of several days, or long distances may be covered in short periods (Lyons and Kanehl 2002). For example, a smallmouth bass migrating to downstream wintering habitats in Wisconsin moved 19 km in 24 hours (Langhurst and Schoenike 1990). Latitudinal differences in temperature and regional variation in annual temperatures exert considerable influence on smallmouth bass distribution, abundance, growth, and survival. A model using temperature, food availability, and lake depth to predict young-of-the-year growth and winter mortality accurately delimited the northern distributional limit of the species (Shuter and Post 1990). Average July temperatures 10◦ C) (Beamesderfer and North 1995). In a study of 129 geographically widespread populations, temperature-related climate differences were significantly related to growth and were most influential in the first 4 years of life (Dunlop and Shuter 2006). On a regional scale, population structure of smallmouth bass in the Laurentian Great Lakes closely tracked changes in water temperatures over several decades. Notably, steep declines in growth and year-class strength occurred with minor temperature shifts (mean shifts 10◦ C before spawning) than small mature males; females show similar size-related timing in spawning (Ridgway et al . 1991b; Wiegmann et al . 1992; Baylis et al . 1993; Lukas and Orth 1995). Smallmouth bass from the Tennessee River exposed to water temperatures of 2.6, 5.2, and 8.0◦ C above ambient temperature (beginning in December) showed spawning peaks of 9, 16, and 25 days, respectively, before control fish exposed to ambient river water temperatures (Wrenn 1984). Likewise, in a thermally unstable, but heated effluent canal in Lake Erie, spawning of smallmouth bass was advanced about 1 month relative to spawning in the lake (Cooke et al . 2003a). Simulated, compressed winter conditions (short photoperiods, temperatures ∼ 6◦ C) followed by
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Centrarchid fishes
20 to 22 days of exposure to increasing photoperiod (14 hours) and temperature (18◦ C) induces out-of-season spawning, but increasing temperature alone does not appear to induce spawning (Cantin and Bromage 1991). Male smallmouth bass establish a territory and use caudal sweeping to excavate a circular depressional nest down to coarse gravel–cobble substrates, bedrock, or even hard clay. Nests average 45 to 93 cm in diameter and are often near (or just downstream of) rocky or woody cover. In lakes and reservoirs, nests are usually placed in water 2 hours and involve 103 female shudders at 30- to 60-second intervals with up to 50 eggs released per shudder. On completion of the bout, the male drives the female from the nest (Reighard 1906; Schneider 1971; Neves 1975). Multiple complete spawning observations, female batch fecundity, and egg developmental stages in nests in natural settings indicate that most males mate with one female, but some males may mate sequentially (or simultaneously) with more than one female (Beeman 1924; Hubbs and Bailey 1938; Neves 1975; Vogele 1981; Ridgway et al . 1989; Wiegmann et al . 1992). Large guardian males are more likely to successfully attract and spawn with females, but in some populations, many males of various sizes build nests but are unsuccessful in attracting mates (Winemiller and Taylor 1982; Wiegmann et al . 1992; Baylis et al . 1993). Of males spawning with females, large guardian males receive more eggs and defend the brood more tenaciously than small guardian males, ultimately producing larger broods, which may in part explain the apparent female mate preference for larger males (Neves 1975; Ridgway and Friesen 1992; Lukas and Orth 1995; Wiegmann and Baylis 1995; Wiegmann et al . 1992, 1997; Knotek and Orth 1998). Mature ovarian eggs average from 1.60 to 2.75 mm diameter, and fertilized, water-hardened eggs from 2.0 to 3.5 mm diameter (Meyer 1970; Smitherman and Ramsey 1972; Hubert 1976; Vogele 1981; Wrenn 1984; Cooke et al . 2006). Fecundity increases with female weight, length, and age (Clady 1975; Hubert 1976; Kilambi et al . 1977; Vogele 1981; Serns 1984; Dunlop et al . 2005b). Bimodal egg size classes occur in ovaries of spawning-ready females, suggesting that females have the potential to spawn multiple batches of eggs in a single spawning season. However, over the relatively short
Centrarchid identification and natural history
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spawning season secondary stage ova do not appear to mature after the initial batch is spawned, being resorbed in summer (Hubert and Mitchell 1979; Vogele 1981). The relationship between potential batch fecundity (Y) and total weight or length (X) are described by the linear functions, Y = −1, 347 + 13.65X, where X is weight in grams (n = 21, R2 = 0.85), or Y = −1225.15 + 59.39X, where X is TL (n = 74, R2 = 0.39) (formulas from Vogele 1981 and Raffetto et al . 1990, respectively; see also, Hubert 1976; Kilambi et al . 1977; Dunlop et al . 2005b). At 549 g (about 335 mm TL), a female can potentially produce 6147 mature eggs in a single batch (range: 1724 eggs at 221 g to 21,467 eggs at 1471 g). Average number of eggs per nest ranges from 2149 to 7757 (>19, 000 in some nests) (Pflieger 1966a; Clady 1975; Neves 1975; Vogele 1981; Raffetto et al . 1990; Wiegmann et al . 1992). The adhesive, grayish white to pale yellow fertilized eggs hatch in 6.4 days at 16◦ C (2.4 days at 22◦ C, from formula in Shuter et al . 1980). Larvae are 4.4 to 6.8 mm TL at hatching, and depending on water temperature, are free swimming at a size of 8.1 to 10.1 mm TL in 4 to 16 days after hatching (Reighard 1906; Beeman 1924; Tester 1930; Hubbs and Bailey 1938; Meyer 1970; Hardy 1978; Shuter et al . 1980; Vogele 1981; Wrenn 1984; Ridgway and Friesen 1992). At swim-up, smallmouth bass fry begin a diel cycle of moving away from the nest at dawn and returning to the nest at dusk, and the guardian male shows parallel behavior (Ridgway 1988). During the swim-up phase, the brood disperses over about 13.4 m2 relative to the guardian male’s nest range of 22.7 m2 . Later, during the juvenile guarding phase, the brood disperses in the day time over 82.4 m2 , and the male over 176.9 m2 . At dusk, fry and male ranges decrease to 3.1 and 20.7 m2 , respectively. The male apparently responds to changes in brood dispersal and not vice versa, because the diurnal contraction and expansion of the brood continues when males are removed (Scott et al . 1997). Juvenile smallmouth bass show nest-site fidelity. In an Ontario lake, age-0 smallmouth bass dispersed little beyond 200 m of their nest of origin by fall, a time long after parental males ceased brood guarding (Gross and Kapuscinski 1997; Ridgway et al . 2002). Likewise, stream-dwelling age-0 smallmouth bass appear to remain near the spawning areas for the first summer of life (Lyons and Kanehl 2002). Male smallmouth bass guard and vigorously defend the nest, eggs, and larvae 24 h/d for 2 to 7 or more weeks, depending in part on male size and energy reserves, spawning time, and water temperatures (e.g., Pflieger 1966a; Neves 1975; Vogele 1981; Hinch and Collins 1991; Ridgway and Friesen 1992; Scott et al . 1997; Knotek and Orth 1998; Cooke et al . 2002a; Cooke et al . 2006). Over eight nesting seasons in a northern lake, average duration of male parental care ranged from 9.4 to 16.4 days (up to 21 days) before swim-up and 9.2 to 11.8 days after swim-up (up to 27 days) (Ridgway and Friesen 1992). Male defense behaviors and swimming activity increase as the offspring progress from egg to hatching, peak before swim-up, and begin to decrease after swim-up (Ridgway 1988; Ongarato and Snucins 1993; Cooke et al . 2002a). Nevertheless, males shift from active and close defense of a brood confined to the nest before swim-up to more distant but vigilant patrolling of dispersed larvae and juveniles (Scott et al . 1997). Guardian male feeding is curtailed or at least dramatically reduced, which in turn reduces and perhaps depletes energy reserves (Hinch and Collins 1991; Gillooly and Baylis 1999; Mackereth et al . 1999; Cooke et al . 2002a; Steinhart et al . 2005). Large males show higher intensity and longer duration of offspring defense; small guardian males can abandon the brood early or may show little or no defense of juveniles, perhaps as a result of reduced or depleted energy reserves (Ridgway and Friesen 1992; Philipp et al . 1997; Mackereth et al . 1999). Males experiencing brood loss from simulated predation also show less nest defense and are more likely to completely abandon the brood (Philipp et al . 1997; Suski et al . 2003). Compelling evidence of an alternating life history strategy is documented for a smallmouth bass population in Nebish Lake, Wisconsin. Unlike the alternative reproductive strategy of cuckoldry seen in some male Lepomis, successive generations of male smallmouth bass in this population alternate their age at first reproduction between ages 3 and 4 (Raffetto et al . 1990; et al .Wiegmann et al . 1992, 1997; Baylis et al . 1993). Micropterus males are typically iteroparous (reproducing in multiple years), but males in this closed population are essentially semelparous (reproducing once in a lifetime). Reproduction can begin at age 3, but the life history decision for time of first reproduction is conditional on male size at age 3, with large age-3 males being likely to reproduce, and small age-3 males being likely to delay reproduction until age 4 or older. In turn, size at age 3 is determined largely in early ontogeny and is likely a function of birth date. Large, older males (age 4 or older) spawn earlier (average about 4–5 days) in the spring than mature, spawning age-3 males. The late spawning, age-3 males are more likely to produce a cohort of small age-3 males that in turn are more likely to delay reproduction until age 4 or older. Conversely, small age-3 males that delay reproduction until age 4 (or older) are more likely to produce a cohort of large, reproductively active age-3 males. Hence, an alternation of time to maturation is sustained over multiple years and appears to be mediated by just a few days difference in birth date (Baylis et al . 1993; Wiegmann et al . 1997). Nest associates: Longnose gar, Lepisosteus osseus (Goff 1984); common shiner, Luxilus cornutus (Hunter and Wisby 1961); orangethroat darter, Etheostoma spectabile (Pflieger 1966b).
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Centrarchid fishes
Freshwater mussel host: Confirmed host to A. ligamentina, L. cardium, L. fasciola, L. higginsii, L. radiata, L. rafinesqueana, L. reeviana, L. siliquoidea, and V. iris (Coker et al . 1921; Zale and Neves 1982; Waller and Holland-Bartels 1988; Barnhart and Roberts 1997; O’Dee and Watters 2000). Putative host to Lampsilis abrupta and Lexingtonia dolabelloides (unpublished sources in OSUDM 2006). Conservation status: The smallmouth bass is secure throughout its range, but native populations in Kansas, along the western periphery of the natural range, are considered vulnerable (NatureServe 2006). Similar species: Spotted bass have a black midlateral stripe (no vertical bars) and rows of black spots along the lower sides; redeye bass have white or orange edges on the caudal fin lobes and rows of black spots along the lower sides; Florida bass and largemouth bass have a dark, midlateral stripe, a deep notch between the soft and spiny dorsal fins, and in adults, the mouth reaches beyond the rear margin of the eye (Page and Burr 1991). Systematic notes: Micropterus dolomieu and M. punctulatus form a sister pair, which is basal to all other Micropterus (Kassler et al . 2002; Near et al . 2003, 2004, 2005). Morphological taxonomists traditionally related M. dolomieu to M. coosae (Hubbs and Bailey 1940; Ramsey 1975). Although only two subspecies of M. dolomieu are usually recognized, the species as currently conceived appears to consist of several distinct evolutionary lineages. The subspecies M. d. velox was described from tributaries of the Arkansas River in southwestern Missouri, northeastern Oklahoma, and northwestern Arkansas based on color, body shape, and modal differences in dorsal ray counts (Hubbs and Bailey 1940). Intergrade populations between M. d. dolomieu and M. d. velox were considered tentatively to occupy the remainder of the southern Ozark and Ouachita uplands, exclusive of the lower Missouri River, and M. d. dolomieu the remainder of the range. Limited sampling of mitochondrial and nuclear DNA sequences did not detect geographic differences among M. dolomieu populations (Kassler et al . 2002; Near et al . 2003, 2004), but nuclear-encoded allozyme loci provide evidence for significant genetic substructuring in the Ozark and Ouachita uplands (Stark and Echelle 1998). Three different clades of M. dolomieu inhabiting the Ozark and Ouachita uplands are evident: (1) the Ouachita smallmouth bass in the Little and Ouachita river drainages; (2) the Neosho smallmouth bass from the southwestern Ozarks in the Neosho and Illinois rivers and smaller tributaries of the middle Arkansas River; and (3) a clade comprising all other populations on the Ozark Plateau (White, Black, St. Francis, Meramec, and Missouri rivers). The latter clade was similar genetically to populations from the upper Mississippi and Ohio River basins (Stark and Echelle 1998). Importance to humans: The smallmouth bass is rivaled only by the Florida bass and the largemouth bass as the most sought-after and valued species in the black bass recreational fishery. Until at least 1932, tons of smallmouth bass were taken commercially by hook and line and by net in Canada, until the species was restricted as a noncommercial sport fish (Scott and Crossman 1973). The smallmouth bass reaches a relatively large size, is an intense, strong fighter when hooked, and over its broad distribution flourishes in high-quality lakes, reservoirs, and upland rivers and streams, all attractive attributes to recreational anglers. As a primary North American recreational fish, the smallmouth bass is the focus of intense fisheries research and management efforts increasingly aimed at maintaining quality- and trophy-size catches for anglers (e.g., Reed et al . 1991; Beamesderfer and North 1995; Kubacki et al . 2002; Noble 2002). Not unexpectedly, techniques for catching smallmouth bass are the subject of a continuous stream of media from the recreational fishing industry (e.g., magazine articles, books, videos). Like other black bass the species is taken by a number of methods including dry flies, wet flies, popping bugs, lures, spinners, jigs, and plastic worms. Effective natural baits include leeches, soft crayfish, hellgrammites, minnow-tipped jigs, frogs, and salamanders. Although most often taken in lakes and reservoirs, smallmouth bass anglers, particularly a growing contingent of fly fishers seeking a quality fishing experience, wade or fish from small boats and canoes in scenic upland streams and rivers (Becker 1983; Etnier and Starnes 1993; Pflieger 1997). The flesh is white, firm, and flaky with fine flavor, being regarded by gourmets as superior table fare (Becker 1983).
13.9.4 Micropterus floridanus Lesueur 13.9.4.1 Florida bass Characteristics: See generic account for general characteristics. Elongate, slightly compressed body, depth about 0.24 to 0.29 of TL, increasing with size. Mouth large, terminal, lower jaw slightly projecting, upper jaw extends beyond
Centrarchid identification and natural history
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posterior edge of eye in adults. Outline of spinous dorsal fin sharply angular. Juncture of soft and spiny dorsal fins deeply emarginate, almost separate. Shortest dorsal spine at emargination of fin, usually 0.3 to 0.4 times the length of longest spine, membranes between short spines deeply incised. Dorsal soft rays, usually 13, 12 to 14; anal soft rays, usually 11, 10 to 12. Gill rakers, 6 to 9. Scales average smaller than largemouth bass. Lateral scales, (65)69 to 73(76); rows above lateral line, (7)8 to 9(10); rows below lateral line, (15)17 to 18(21); cheek scale rows, (10)11 to 13(14); caudal peduncle scale rows, (27)28 to 31(33); pectoral rays, 14 to 15(16). No small splintlike scales on interradial membranes at anal and second dorsal fin bases. Pyloric caeca branched at bases, 26 to 43 or more. Tooth patch absent (rarely a few teeth) on glossohyal (tongue) bone (Bailey and Hubbs 1949; Buchanan 1973; Chew 1974; Ramsey 1975; Kassler et al . 2002). Size and age: Size at age 1 ranges from 142 to 310 mm TL for males and 116 to 330 mm TL for females (Allen et al . 2002). Age and weights of trophy Florida bass (n = 810, ≥4.5 kg) obtained from taxidermists across Florida revealed a maximum age of 16 (average 9.7 years), a maximum weight of 7.9 kg (average 5.0 kg), and a maximum length of 762 mm TL (average 661 mm) (Crawford et al . 2002). Florida state record, 7.85 kg (FFWCC 2006). Females grow faster and live longer than males; nearly all large individuals of Florida bass (>400 mm TL) are females (Allen et al . 2002; Crawford et al . 2002; Bonvechio et al . 2005; all cited studies include a few likely populations of M. floridanus × M. salmoides intergrades in northern Florida). Coloration: Broad dark olive to olive black, midlateral stripe on caudal peduncle becoming disrupted anteriorly into a series of more or less distinct blotches, the midlateral stripe often faint in large adults. Silver to brassy green above (brownish in tea-stained water) with dark olive mottling. Scattered dark specks on lower sides; whitish below. Iris brown. Young (300 mm TL) feed about equally on fish (e.g., other centrarchids, clupeids, anchovies, topminnows, lake chubsuckers, silversides, minnows, darters) and decapods (crayfish and grass shrimp, if available) (McLane 1948, 1950; Chew 1974; Schramm and Maceina 1986; Huskey and Turingan 2001; Crawford et al . 2002). Young-of-the-year (13–30 mm TL) feed heavily on cladocerans, copepods, amphipods, and aquatic insects but with growth (31–75 mm TL) cease zooplankton use and begin including higher volumes of grass shrimp and fish (e.g., mosquitofish, silversides, topminnows). By 75 mm TL, fish and decapods constitute most of the diet biomass (Carr 1942; Chew 1974; Huskey and Turingan 2001; Allen and Tugend 2002). Florida bass feed by using combinations of ram (i.e. rapid acceleration of the body) and suction (i.e. rapid expansion of buccal cavity) strike modes on prey (Sass and Motta 2002). Feeding activity appears to occur randomly during the day (Chew 1974), and in captivity, Florida bass digestion rates are rapid (relative to warmouth, L. gulosus), and individuals feed voraciously even when considerable food from previous meals remains in the stomach (Hunt 1960). In the St. Johns River, Florida, early naturalists reported groups of hundreds to thousands of Florida bass pursuing and feeding on enormous schools of threadfin shad. Attacks by the bass on the shad resulted in the surface boiling with activity for several minutes at a time (McLane 1948). Focal animal observations on Florida bass (300 TL) hunted only with groups of other bass, but small individuals (0.6 times length of longest spine. Dorsal soft rays, 12 to 13; anal soft rays, 10 to 11. Gill rakers, usually 5. Relatively large scales. Lateral scales, 57 to 65; rows above lateral line, 6 to 9; rows below lateral line, 14 to 19; cheek scale rows, 9 to 15; caudal peduncle scale rows, 27 to 31; pectoral rays, (15)16(17). Small splintlike scales on interradial membranes at anal and second dorsal fin bases (>60 mm SL). Pyloric caeca single, rarely branched, 10 to 13. Tooth patch on glossohyal (tongue) bone (Bailey and Hubbs 1949; Ramsey and Smitherman 1972; Page and Burr 1991; Kassler et al . 2002). Size and age: Size at age 1 ranges from 146 to 206 mm TL. Large individuals are >305 mm TL, weigh 400 g, and reach age 7+ (maximum 402 mm TL and age 9+ for males, age 12+ for females) (Bass and Hitt 1973; Page and Burr 1991; Cailteux et al . 2002; Bonvechio et al . 2005). World angling record, 1.75 kg, Florida (IGFA 2006). Females grow faster and live longer than males, and in a given population, 60% to 100% of individuals >305 mm TL are females (Bonvechio et al . 2005). Coloration: Color similar to M. salmoides but usually brown overall, and sides marked with about 12 vertically elongate, lateral blotches. Blotches anteriorly are much wider than their interspaces, becoming more confluent with age. The blotches fuse on the caudal peduncle to form a relatively uniform, wide lateral band. Ventrolateral longitudinal streaks are weakly developed. Iris red. Young with a series of thin, closely spaced vertical bars along the sides of the body. Cheeks, breast, and lower sides colored brilliant turquoise blue in nesting males, less so in non-nesting individuals (Bailey and Hubbs 1949; Gilbert 1978; Page and Burr 1991). Native range: The Suwannee bass is native to the Suwannee and Ochlockonee Rivers, Florida and Georgia (MacCrimmon and Robbins 1975; Page and Burr 1991). The provenance of populations in the Wacissa (Aucilla River drainage), Wakulla, and St. Marks rivers of Florida is uncertain (Koppelman and Garrett 2002; Cailteux et al . 2002; Bonvechio et al . 2005) but, given the lack of historical records, are likely introduced. Electrofishing catch data indicate that the species is most abundant in the Wacissa River (Aucilla River drainage) and Santa Fe River (Suwannee River drainage) (Schramm and Maceina 1986; Cailteux et al . 2002; Bonvechio et al . 2005). Habitat: The Suwannee bass occurs in a variety of habitats in cool, clear, spring-fed rivers, which characteristically have limestone substrates (often covered with sand); alkaline, hard water; relatively stable thermal regimes; and dense submersed macrophyte beds (Bass and Hitt 1973; Gilbert 1978; Schramm and Maceina 1986; Cailteux et al . 2002). In the Santa Fe River, individuals (>150 mm TL) are associated with fallen trees over sandy substrate; shallow bedrock riffles (0.7–3.0 m deep); vegetated (eelgrass), gravel–sand riffles; deep vertical rock drop-offs (to 3 m); and shallow, sandy, gently sloping vegetated banks (0.5–1.0 m deep). Small individuals are most common around fallen trees but occur in a variety of flowing and nonflowing habitats (Schramm and Maceina 1986). Individuals also occupy spring runs of river tributaries where they seek cover under dense overhanging or floating vegetation (Gilbert 1978). Food: The Suwannee bass is a top carnivore, extensively exploiting crayfishes for food. Crayfishes are the predominant food of individuals >150 mm TL, and for large fish (>300 mm TL), the diet is almost exclusively crayfishes. Fish rank second and freshwater shrimp third in importance in the diet; other crustacea, such as blue crabs, and a few aquatic insect larvae are also consumed. Juveniles (167 mm TL) to consume larger prey items at a given size than the sympatric congener. Stomach contents of 142 Suwannee bass sampled in daylight hours from May to August revealed no obvious feeding periodicity (Schramm and Maceina 1986).
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Reproduction: Size and age at maturity are not well documented, and little is published on reproductive behavior and biology of this unique, range-restricted Micropterus. Gonads of the sexes are distinguishable at minimum sizes of 125 mm SL in males and 142 mm SL in females, but the smallest females reported with mature ova are ≥215 mm SL (Bass and Hitt 1973). On the basis of female reproductive condition and other observations, spawning apparently begins in February or March as water temperatures reach 18 to 20◦ C and continues into June. Females with ripe ova are taken from February to May, spent females begin to appear in April with the largest numbers occurring in May. Suwannee bass nests in rivers have been noted in April, and spawning occurred in experimental ponds in Alabama in early April (Bailey and Hubbs 1949; Hellier 1967; Smitherman and Ramsey 1972; Bass and Hitt 1973). Young 60 mm SL). Pyloric caeca, single, rarely branched, 10 to 13. Tooth patch present on glossohyal (tongue) bone (Hubbs 1927; Hubbs and Bailey 1940, 1942; Applegate 1966; Bryan 1969; Ramsey and Smitherman 1972; Williams and Burgess 1999). Size and age: Size at age 1 averages about 113 mm TL but varies considerably among habitats and across the geographic range (population averages range from 66 to 216 mm TL) (Vogele 1975b; Webb and Reeves 1975; Carlander 1977; Olmsted and Kilambi 1978; DiCenzo et al . 1995; Pflieger 1997; Maceina and Bayne 2001). Growth rate trends higher in reservoirs than in streams (Vogele 1975b), and the Alabama spotted bass, M. p. henshalli, lives longer and reaches a larger size than the northern subspecies, M. p. punctulatus (DiCenzo et al . 1995). However, the Alabama spotted bass may represent a distinct taxon and perhaps be only distantly related to M. punctulatus (e.g., Kassler et al . 2002). Few individuals exceed 425 mm TL, 2.0 kg, and ages 6+ (maximum about 640 mm TL and age 11+) (Gilbert 1973; Webb and Reeves 1975; Carlander 1977; Olmsted and Kilambi 1978; Page and Burr 1991; DiCenzo et al . 1995; Wiens et al . 1996; Maceina and Bayne 2001). World angling record, 4.65 kg, California (IGFA 2006). Females of the Alabama spotted bass, M. p. henshalli, and perhaps other spotted bass populations (e.g., Ryan et al . 1970), can live longer than males (age 8+ versus age 5+) and after the third year show faster growth and weigh more than males (Webb and Reeves 1975). Coloration: Rows of small black spots on yellow-white lower sides form horizontal lines. Dark midlateral stripe or series of partly joined blotches along light olive to yellowish green side. Caudal spot dark, darkest on young. Light green-gold dorsally with dark olive, often diamond-shaped mottlings. Young (95% of remaining M. dolomieu mtDNA haplotypes (and nuclear alleles) in the lake population were found in fishes of hybrid ancestry between the introduced and native Micropterus. Similar patterns indicative of introgressive swamping occurred when M. punctulatus was introduced into a native population of M. dolomieu in South Moreau Creek (Missouri River drainage), Missouri (Koppelman 1994), and are suggested for introductions of M. p. henshalli into a native population of M. coosae in Keowee Reservoir (Savannah River drainage), South Carolina (Barwick et al . 2006). Habitat: The spotted bass inhabits gravelly flowing pools and runs of creeks and small to medium rivers and reservoirs (Page and Burr 1991). In streams, spotted bass are commonly associated with low-velocity pools, particularly those with vegetation, log complexes, rootwads, or undercut banks (Lobb and Orth 1991; Scott and Angermeier 1998; Tillma et al . 1998; Horton and Guy 2002; Horton et al . 2004). The habitat requirements of the species can be broadly characterized as intermediate between those of the smallmouth bass and largemouth bass. The spotted bass is associated with warmer, more turbid water than smallmouth bass, and faster, less productive waters than the largemouth bass (Trautman 1981; Layher et al . 1987; Pflieger 1997). Nevertheless, spotted bass frequently co-occur with largemouth bass, smallmouth bass, and redeye bass but generally show some spatial segregation from co-occurring Micropterus, in cover type,
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longitudinal distribution, or water depth (e.g., Viosca 1931; Vogele 1975b; Trautman 1981; Buynak et al . 1989; Matthews et al . 1992; Pflieger 1997; Scott and Angermeier 1998; Sammons and Bettoli 1999; Long and Fisher 2005). For example, spotted bass were widely distributed in a Virginia impoundment, but occurred most commonly in areas with fine substrate and woody debris, undercut banks, and bank vegetation as cover, avoiding the steep drop-offs and rocky shorelines frequented by smallmouth bass (Scott and Angermeier 1998). In southern US reservoirs, spotted bass are most abundant in oligo-mesotrophic reservoirs or oligo-mesotrophic reaches of reservoirs with abundance decreasing as eutrophication increases; an opposite pattern occurs for largemouth bass abundance (Buynak et al . 1989; Greene and Maceina 2000; Maceina and Bayne 2001). Although spotted bass may enter relatively high-salinity coastal environments (≤10 ppt), they infrequently occur in coastal marshes with salinities >4 ppt (Peterson 1988, 1991; Peterson and Ross 1991). Relatively little is known about movements of spotted bass. In some populations, indirect evidence suggests massive upstream movement in spring from reservoirs and rivers into tributaries to spawn, followed by a gradual downstream drift of most adults and young to overwinter in large, lower-gradient habitats (Vogele 1975b; Trautman 1981). The average home activity area of radio-tagged spotted bass tracked over multiple seasons in a Kansas stream was 0.39 ha (range, 0.06–1.2 ha). Activity area was correlated positively with body size, and activity areas of up to six fish showed simultaneous overlap. During summer and winter, fish typically remained in one pool, but during spring and fall, fish crossed riffles and moved among pools (Horton and Guy 2002). Food: The spotted bass is an opportunistic carnivore, exploiting prey from the bottom to the water’s surface. The adult diet is dominated in biomass by crayfish if present, fish (e.g., clupeids, darters, minnows, catfishes), and to a lesser extent, immature aquatic insects (Applegate et al . 1967; Gilbert 1973; Vogele 1975b; Scott and Angermeier 1998). Depending on prey availability, consumption of large numbers and volumes of immature aquatic insects may continue up to 150 mm TL or larger. Spotted bass may exploit relatively large numbers and volumes of terrestrial insects (e.g., hymenoptera, beetles, flies, adult odonates) (Smith and Page 1969; Ryan et al . 1970; Vogele 1975a; Scott and Angermeier 1998). The young initially depend on zooplankton (cladocerans and copepods) with juveniles transitioning from large immature aquatic (e.g., mayflies, diptera) insects to fish and crayfish at 50 to 100 mm TL (Applegate et al . 1967; Clady and Luker 1982; Matthews et al . 1992; Scott and Angermeier 1998). Spotted bass are relatively inactive at night, staying close to cover, but move frequently throughout the day (Horton et al . 2004). Even so, diet data reveal no clear diel feeding patterns except for an increase in terrestrial insects in the diet during the day (Scott and Angermeier 1998). Reproduction: Maturity can be reached as early as age 1+ in fast-growing populations, but most individuals do not mature until age 2+ to 3+ (Gilbert 1973; Olmsted 1974; Vogele 1975a,b). Depending in part on latitude and water temperature, spawning occurs over a 1- to 2-month period from March to May or early June, with the most intensive nesting occurring within about 2 weeks of initial spawning activity (Ryan et al . 1970; Gilbert 1973; Olmsted 1974; Vogele 1975a; Sammons et al . 1999; Greene and Maceina 2000). Active nests have been observed at temperatures as low as 12.8◦ C, but most spawning occurs between 14◦ C and 23◦ C (Howland 1932a; Ryan et al . 1970; Smitherman and Ramsey 1972; Gilbert 1973; Olmsted 1974; Vogele 1975a,b; Aasen and Henry 1981; Sammons et al . 1999). The male excavates a solitary, depressional, roughly circular nest by caudal sweeping and removing material with his mouth (Breder and Rosen 1966); nests are spaced widely with densities ranging from 0.5 to 11.3/100 m of shoreline. Most but not all nests are located near cover (e.g., rock overhangs, stumps, submerged tree bases) (Vogele 1975a; Vogele and Rainwater 1975). Nests are 38 to 76 cm in diameter, are located at average water depths of 2.3 to 3.7 m (range, 0.9–6.7 m), and are usually swept out over hard substrates (e.g., sand and gravel, solid rock ledges, flat rocks), but compacted soil and exposed root hairs of flooded trees are also used (Vogele 1975a,b; Aasen and Henry 1981). Males may excavate and defend one to four nest sites for up to 3 days before egg deposition. Limited evidence from tagged males suggests year-to-year fidelity to specific nesting areas (Vogele 1975a). Courtship and spawning are generally typical of other Micropterus, but published documentation is not extensive (e.g., male guiding of female, paired circling) (Miller 1975; Vogele 1975a,b, citing Howland 1932b). Once a female is attracted to the nest, the male guides her in circles about the nest (female inside, male outside), repeatedly biting at her opercle and vent. During courtship, the midlateral stripe in the female disappears (Miller 1975). Courtship behaviors continue for 20 minutes to 1 hour before egg deposition begins. Ultimately, the female deposits eggs (for 1.5 to 5 seconds) by tilting on her side, and the male releases milt in an upright position as is typical for most centrarchids. Courtship and spawning sequences between pairs may require up to 3.5 hours for completion (Vogele 1975a). Most spawning observations involved a single male and female. After spawning, males immediately begin fanning the eggs and continue defending the eggs from numerous, persistent Lepomis and other predators (Vogele 1975a). Mature ovarian eggs range from 1.30 to 2.20 mm diameter (Gilbert
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1973; Vogele 1975a) and fertilized, water-hardened eggs range from 1.60 to 2.30 mm diameter (Smitherman and Ramsey 1972; Vogele 1975a). Fecundity increases with female size. The relationship between potential batch fecundity (Y) and total length (X) is described by the function, log10 Y = −8.222 + 4.779 log10 X(n = 48, R2 = 0.71, data from Olmsted 1974 and Vogele 1975a). At 347 mm TL, a female can potentially produce 8284 mature eggs in a single batch (range: 1728 eggs at 250 mm TL to 26,906 eggs at 444 mm TL, respectively). The adhesive, fertilized eggs hatch in 5 days at 14.4◦ C to 15.6◦ C (Vogele 1975a). Larvae are free swimming at 6.0 to 7.5 mm TL in 4 days and 8 days after hatching at 25◦ C and 15 to 18◦ C, respectively (Vogele 1975a; DiCenzo and Bettoli 1995). Fry emerging from the nest form compact schools that are guarded by the parental male for up to 4 weeks. Schools with fry from different nests may merge into a single large school and be guarded by two parental males. The schools break up as fry reach about 30 mm TL (Vogele 1975a). In hatchery ponds, males apparently exhibited less parental care, abandoning the fry shortly after swim-up (Smitherman and Ramsey 1972; Vogele 1975b). Nest associates: None known. Freshwater mussel host: Confirmed host to L. altilis, L. perovalis, Lampsilis subangulata, V. iris, V. nebulosa, and V. vibex (Neves et al . 1985; Haag and Warren 1997; Haag et al . 1999; O’Brien and Brim Box 1999). Putative host to L. abrupta (unpublished sources in OSUDM 2006). Conservation status: The spotted bass is secure throughout its range, but peripheral populations in Illinois are considered vulnerable (Warren et al . 2000; NatureServe 2006). Lack of resolution of the genetic relationships among populations now regarded as M. punctulatus is of primary conservation concern (Kassler et al . 2002; see section on systematic notes). Similar species: Shoal bass has dark vertically elongate bars on sides and lacks patch of teeth on tongue; redeye bass has white to orange upper and lower edges on caudal fin lobes and young has red medial fins; largemouth bass and Florida bass lack rows of black spots on lower sides and have a deep notch between spiny and soft dorsal fins; young of these species have a bicolored caudal fin (white, black edge); smallmouth bass lacks a distinct lateral stripe (Page and Burr 1991). Systematic notes: Micropterus punctulatus and M. dolomieu form a sister pair that is basal to all other Micropterus (Kassler et al . 2002; Near et al . 2003, 2004, 2005). As currently conceived, the long-presumed polytypy of M. punctulatus (Hubbs and Bailey 1940) appears to subsume two relatively distantly related and divergent species of Micropterus. Morphological and genetic data indicate that a small-scaled form, the Alabama spotted bass (nominal M. p. henshalli ), occurs in Mobile Basin (Hubbs and Bailey 1940; Gilbert 1973; Kassler et al . 2002). Although intergrades between M. p. punctulatus and M. p. henshalli were suggested from limited samples from west of Mobile Basin to the Lake Pontchartrain system (Hubbs and Bailey 1940), more extensive meristic data revealed no evidence of intergradation in that region (Gilbert 1973). However, individuals above the Fall Line in Mobile Basin were assigned to M. p. henshalli and those below the Fall Line were interpreted as intergrades between M. p. henshalli and M. p. punctulatus (Gilbert 1973). The putative intergrades could just as easily represent in situ differentiation of quasi-isolated populations of Alabama spotted bass, rather than intergradation. Importantly, mitochondrial DNA analyses from limited population sampling indicate that the form in Mobile Basin is highly divergent from M. p. punctulatus (e.g., fixed allelic differences at multiple gene loci, fixed haplotype differences, sequence divergence of 10.3%) and is genetically most similar to M. coosae (Kassler et al . 2002). Unfortunately, M. p. henshalli has been introduced outside the native range in Mobile Basin and has introgressed with native Micropterus (Pierce and Van Den Avyle 1997). The resolution of the relationships of the Alabama spotted bass to other Micropterus awaits a thorough genetic analysis across populations in the Mobile Basin. The subspecies M. p. wichitae, ostensibly restricted to a single stream in the Red River drainage, Oklahoma (Hubbs and Bailey 1940), was based on M. punctulatus × M. dolomieu hybrids and is not valid (Cofer 1995). The subspecies M. p. punctulatus occupies the remainder of the range (Gilbert 1973). Importance to humans: Ecologically, the spotted bass can function as the only top carnivore in small, even intermittent, headwater streams and is often the dominant top predator in large rivers and reservoirs (Cross 1967; Trautman 1981; Pflieger 1997). The spotted bass is also a popular sport fish in streams and reservoirs throughout the southeastern United States. The species is sought in streams by anglers favoring fly fishing or ultralight tackle (Cross 1967; Ross 2001). The largest spotted bass are taken in reservoirs and spillways where food availability is higher than in most streams (Ross
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2001). In southern US reservoirs, spotted bass can be the dominant or co-dominant Micropterus and constitutes a sizable proportion of the black bass catch (e.g., 60%) and harvest (e.g., 50%) (Webb and Reeves 1975; Novinger 1987; Buynak et al . 1989, 1991; DiCenzo et al . 1995; Pflieger 1997; Sammons et al . 1999; Sammons and Bettoli 1999; Long and Fisher 2005). The spotted bass often co-occurs with the largemouth bass or smallmouth bass in reservoirs, where most management effort is usually focused on the latter two species (e.g., Maceina and Bayne 2001; Long and Fisher 2005). Because of its slower growth and high abundance in some reservoirs, fishery managers combine liberalized harvest of spotted bass with increased length limits for largemouth bass (or smallmouth bass) to reduce exploitation and to increase the size of the latter (e.g., Buynak et al . 1991; Long and Fisher 2005). The spotted bass takes the same lures (e.g., spinner baits, plastic worms, jigs, crank baits) and live baits (e.g., minnows, crayfishes, salamanders) as other black bass. Anglers consider their strike more aggressive and their fight more spirited than that of the largemouth bass (Ross 2001).
13.9.7 Micropterus salmoides Lac´ep`ede 13.9.7.1 Largemouth bass Characteristics: See generic account for general characteristics. Elongate, slightly compressed body, depth 0.24 to 0.29 of TL, increasing with size. Mouth large, terminal, lower jaw slightly projecting, upper jaw extends beyond posterior edge of eye in adults. Outline of spiny dorsal fin sharply angular. Juncture of soft and spiny dorsal fins deeply emarginate, almost separate. Shortest dorsal spine at emargination of fin, usually 0.3 to 0.4 times length of longest spine, membranes between short spines deeply incised. Dorsal soft rays, usually 13 or 14, 11 to 15; anal soft rays, usually 11 or 12, 10 to 14. Gill rakers, 7 to 9. Lateral scales, (55)58 to 67(72); rows above lateral line, 7 to 8(9); rows below lateral line, 13 to 17; cheek scale rows, 9 to 11(13); caudal peduncle scale rows, (24)26 to 28(30); pectoral rays, (13)14 to 15(17). No small splintlike scales on interradial membranes at anal and second dorsal fin bases. Pyloric caeca branched at base, 12 to 45. Tooth patch usually absent on glossohyal (tongue) bone, but tooth patch present or absent in San Antonio and Nueces rivers, southwest Texas, and present in ≥50% of specimens in the Rio Grande system, Mexico and Texas (Hubbs and Bailey 1940; Bailey and Hubbs 1949; Applegate 1966; Keast and Webb 1966; Buchanan 1973; Chew 1974; Edwards 1980; Kassler et al . 2002). Size and age: Size at age 1 is highly variable among habitats and across latitudes, ranging from 33 to 271 mm TL (median 102 mm TL) (Carlander 1977; McCauley and Kilgour 1990; Beamesderfer and North 1995; Garvey et al . 2003). Critical periods causing differential size, growth, and survival for age-0 cohorts include time of hatching, onset of piscivory, accumulation of lipids in the fall, and the ability to survive predation, starvation, or both over the first winter (DeAngelis and Coutant 1982; Gutreuter and Anderson 1985; Miranda and Hubbard 1994a,b; Ludsin and DeVries 1997; Maceina and Bettoli 1998; Garvey et al . 1998; Post et al . 1998; Fullerton et al . 2000; Garvey et al . 2000, 2002; see section on habitat). Large individuals can exceed 550 mm TL, weigh >3.5 kg, and attain age 8+ to 15+ (Carlander 1977; Beamesderfer and North 1995). The oldest largemouth bass and longest-lived Micropterus is a 23- or 24-year-old individual (584 mm TL) from New York (Green and Heidinger 1994). The world angling record for all Micropterus (and all centrarchids) is a largemouth bass weighing 10.1 kg (∼ 787 mm TL) that was caught in Georgia in 1932 (IGFA 2006). At least in some populations, older females (age 4+) are longer than males, and most older individuals are females (Webb and Reeves 1975; Carlander 1977). Coloration: Broad olive or olive black midlateral stripe formed of confluent or nearly confluent blotches. Silver to brassy green (brownish in tea-stained water) above with dark olive mottling. Scattered dark specks on lower sides; whitish below. Iris brown. Young (10◦ C and latitude), and instantaneous natural mortality rate was correlated positively with mean air temperature (Beamesderfer and North 1995). Likewise, average length by fall of age-0 largemouth bass is related positively to latitude and presumably temperature (Garvey et al . 2003). Temperature effects are directly or indirectly related to several critical events in the first year of life including hatch date, length of growing season, transition to piscivory, fall lipid accumulation, winter food availability, and the duration and severity of winter (Kramer and Smith 1960a, 1962; Adams et al . 1982a,b; Isely et al . 1987; Miranda and Hubbard 1994a,b; Ludsin and DeVries 1997; Post et al . 1998; Wright et al . 1999; Fullerton et al . 2000; Jackson and Noble 2000; Fuhr et al . 2002; Philipp et al . 2002). For age-0 fish, winter is often a huge survival bottleneck because of complex interactions of winter severity, food availability, and predation. When water temperatures are 80 mm TL) increased with increased bass abundance. Small fishes remaining in bass-containing pools occupied shallow pool margins, but those in pools without bass used the entire pool. Larval minnows and larval Lepomis were only found in pools that
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contained, or had contained, largemouth bass. Experimental manipulation of bass and Lepomis larvae in stream pools indicated that bass presence enhanced short-term survival of the larvae, likely an indirect effect of the shift in small fishes that prey on the larvae (Harvey 1991a). A particularly strong seasonal interaction can occur between largemouth bass, an algae-grazing minnow (Campostoma anomalum), and attached algae in stream pools. Large schools of Campostoma grazing in stream pools can dramatically reduce algal biomass and composition on stream substrates (Power and Matthews 1983; Matthews et al . 1987; Power et al . 1988) and influence the life histories of other invertebrates as well (Vaughn et al . 1993). In a small prairie-margin stream in Oklahoma, largemouth bass (>70 mm SL) and Campostoma showed complementary distributions among stream pools with differential crops of periphyton during summer low flow (Power and Matthews 1983; Power et al . 1985). Pools with bass had lush standing crops of epiphyton covering rocky substrates, but in the Campostoma pools, epiphyton was confined to pool margins, and most rocky substrates were bare. Experimental addition of bass to pools caused Campostoma to immediately emigrate from the pool or move to shallow water margins of the pool. Those that did remain in bass pools spent significantly less time in feeding and more time in cover than they did before bass were added. After bass addition, the standing crop of algae in pools increased significantly within 10 to 13 days (Power et al . 1985). The pattern of abundance of adult largemouth bass and small fishes in streams is congruent with that observed in lake communities. Several studies demonstrate the shift of juvenile bluegill to vegetated or shallow littoral zones as a refuge from predation by Micropterus (e.g., Savino and Stein 1982, 1989a,b; DeVries 1990; Gotceitas 1990b; Gotceitas and Colgan 1990) and others demonstrate the indirect effects of largemouth bass on the zooplankton prey of bluegills or other Lepomis (e.g., Hambright et al . 1986; Werner and Hall 1988; Turner and Mittelbach 1990; Hambright 1994). For example, in pond experiments using largemouth bass and small bluegills, the bass induced a habitat shift in small bluegill, resulting in size distributions skewed toward larger bluegill, a direct predation effect of bass. In turn, the shift to larger bluegill produced pronounced differences in zooplankton abundance and size structure (e.g., three cladocerans and the phantom midge became more abundant in the bass treatment), an indirect effect of bass on the aquatic community (Turner and Mittelbach 1990). A long-term lake study in which largemouth bass were eliminated by a natural event (1978) and then reintroduced (1986) is further illustration of their role as keystone species in some lakes (Mittelbach et al . 1995; see also Carpenter et al . 1987; Hall and Ehlinger 1989; Drenner et al . 2002). Elimination of bass was followed by a dramatic increase in planktivorous fish (e.g., golden shiner, 400,000/lake), the disappearance of large zooplankton, and the appearance of many small-bodied cladocerans, states which were maintained throughout the period of absence of the bass. On reintroduction of largemouth bass, the lake steadily returned to its previous state. Planktivore numbers decreased by two orders of magnitude (golden shiners being practically eliminated), large-bodied zooplankton reappeared and dominated the zooplankton, and the suite of small-bodied cladocerans disappeared. Total zooplankton biomass increased 10-fold and water clarity increased significantly. Reproduction: Maturity is usually reached by age 2+ to 4+ at minimum sizes of about 250 to 300 mm TL but can occur at age 1+ in fast-growing populations or be delayed until age 5+ in cool north temperate waters (Bryant and Houser 1971; Webb and Reeves 1975; Carlander 1977; Becker 1983). Spawning activity can begin in early spring at a water temperature as low as 12◦ C, but most individuals initiate spawning after the water temperature reaches and exceeds 15◦ C. The spawning season extends over 2 to 10 weeks, peaks between water temperatures of 15 and 21◦ C, and winds down as waters warm to and consistently exceed 24◦ C. Spawning occurs from mid-May to mid-June or even early July at north temperate latitudes and shifts to earlier dates at progressively lower latitudes (e.g., mid-March to May or early June in Mississippi and Alabama) (Kramer and Smith 1960a; Allan and Romero 1975; Becker 1983; Miller and Storck 1984; Isely et al . 1987; Goodgame and Miranda 1993; Annett et al . 1996; Post et al . 1998; Sammons et al . 1999; Greene and Maceina 2000; Cooke et al . 2006). Large adult male and female largemouth bass spawn before smaller adults. The earlier hatched young of large bass often gain and maintain a distinct size advantage over the later hatched young of smaller bass, a size advantage that may increase probability of survival to age 1+ (Miller and Storck 1984; Miranda and Muncy 1987; Goodgame and Miranda 1993; Phillips et al . 1995; Ludsin and DeVries 1997; Sammons et al . 1999; Pine et al . 2000). Males use caudal sweeping to excavate circular, depressional nests (0.6–1.0 m diameter) 1 to 2 days before spawning (Kramer and Smith 1962; Cooke et al . 2001b). Males can successfully sweep out nests over a variety of substrates (e.g., silt to boulders, stump tops, logs, clay slabs), but coarse gravel and sand and the roots and stems of aquatic vegetation are substrates most often used (Reighard 1906; Miller and Kramer 1971; Allan and Romero 1975; Annett et al .
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Centrarchid fishes
1996; Hunt et al . 2002). Most males select nest sites near simple cover (e.g., horizontal log, tree trunk) where they suffer less nest intrusion by brood predators and expend less effort in aggressive actions than males selecting sites near complex cover (e.g., brush piles, patches of aquatic macrophytes) (Annett et al . 1996; Hunt et al . 2002). Although a few nests have been reported from >6 m depth, most nests are placed in water 90 mm SL) diet is dominated by small fishes, mostly minnows (e.g., Notropis, Cyprinella, Campostoma) and other centrarchids, but also includes large numbers of mayfly, dragonfly, dipteran, hemipteran, and megalopteran larvae, a few bees and wasps, and an occasional amphibian. Large adults (>150 mm SL) consume relatively large volumes of crayfish. Fish prey associated with flowing water (e.g., blacktail shiner, darters, channel catfish) are taken most often, an indication of the primary foraging habitat of Guadalupe bass. By volume, the diet of young bass (15–30 mm SL) is dominated by mayfly, odonate, and hemipteran larvae. In bass between 30 and 90 mm SL, increasing volumes of fish are consumed, but invertebrates remain important components of the diet of bass 30%) of the angling effort is directed at crappies (e.g., Larson et al . 1991; Reed and Davies 1991; St. John and Black 2004). A growing contingency of crappie anglers are considered “specialists,” similar to many black bass anglers, because they fish year round for crappies to the near exclusion of other species. The relatively recent advent of crappie clubs and fishing tournaments, dubbed crappiethons, are further evidence of the continued and growing popularity of sport fishing for these centrarchids (Larson et al . 1991; Allen and Miranda 1996). Generic characteristics: Deep, extremely compressed body, depth about 0.33 to 0.48 of SL. Long to very long predorsal region with sharp dip over eye in dorsal profile. Dorsal fin base equal to or shorter than distance from center of eye to dorsal fin origin. Head small. Eye large, diameter equal to or slightly greater than snout length. No black teardrop; no black spot in soft dorsal fin. Dorsoposterior margin of opercle shallowly emarginate. Preopercle posterior margin serrate. Long dorsal fin, 6 to 8 spines, 13 to 18 rays, 20 to 24 total; and long anal fin, 5 to 8 spines, 14 to 18 rays, 23 to 24 total. Spiny and soft dorsal and anal fins continuous, smoothly rounded, similar in length, and nearly symmetrical. Emarginate to shallowly forked caudal fin. Rounded pectoral fin. Long, slender gill rakers, 25 to 32. Ctenoid scales. Lateral line complete. Lateral line scales, 34 to 50; cheek scale rows, 5 to 6; branchiostegal rays, 7. Teeth on entopterygoid and glossohyal (tongue, two patches) bones (Bailey 1938; Keast 1968a; Trautman 1981; Becker 1983; Smith 1985; Page and Burr 1991; Etnier and Starnes 1993; Mabee 1993; Jenkins and Burkhead 1994; Smith et al . 1995). Similar species: See account on flier.
13.10.1 Pomoxis annularis Rafinesque 13.10.1.1 White crappie Characteristics: See generic account for general characteristics. Deep, extremely compressed body, depth usually 0.33 to 0.48 of SL. Very long predorsal region with sharp dip over eye in dorsal profile. Dorsal fin base shorter than distance from center of eye to dorsal fin origin. Large, supraterminal, oblique mouth, lower jaw projecting, supramaxilla moderate (≤2 times length of maxilla), upper jaw reaching to or slightly beyond middle of eye. Opercular spot black. Long dorsal fin, (4)5 to 6(8) spines, (12)14 to 15(16) rays; and long anal fin, 6 to 7(8) spines, 16 to 19 rays. Pectoral rays, (14)15(16); vertebrae, 30 to 32(14+18) (Bailey 1938; Trautman 1981; Becker 1983; Page and Burr 1991; Etnier and Starnes 1993; Mabee 1993; Jenkins and Burkhead 1994; Smith et al . 1995). Size and age: Typically reach 131 to 173 mm TL at age 1, but first-year growth is highly variable across latitudes and among habitats (range, 58–310 mm TL, Siefert 1969a; Carlander 1977). Large individuals measure 350 to 400 mm TL, weigh 500 to 800 g, and reach age 6+ to 8+ (maximum 530 mm TL, age 9+) (Carlander 1977; Page and Burr 1991; Etnier and Starnes 1993). World angling record, 2.35 kg, Mississippi (IGFA 2006). Coloration: Gray-green above with silvery blue sides and upper back vaguely barred with about 6 to 10 chainlike double vertical bands (widest at top) as well as dark blotches and green flecks. Chainlike bars and mottling often faint in individuals from turbid water. Whitish to silvery below. Dorsal, anal, and caudal fins with many wavy dark bands and spots. Males become darker during the breeding season (Page and Burr 1991; Etnier and Starnes 1993). Native range: The white crappie is native to the Great Lakes, Hudson Bay (Red River), and Mississippi River basins from New York and southern Ontario west to Minnesota and South Dakota and south to the Gulf of Mexico and in Gulf drainages from Mobile Bay, Georgia and Alabama, west to the Nueces River, Texas (Page and Burr 1991). The species has been introduced and is established over most of the coterminous United States (Fuller et al . 1999).
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Habitat: The white crappie inhabits sand- and mud-bottomed pools and backwaters of creeks and small to large rivers, lakes, ponds, and reservoirs (Page and Burr 1991). The greater adaptability of the white crappie to turbid waters than the black crappie is often noted. Higher relative abundance or success in turbid habitats suggests that the white crappie is more adapted to turbid conditions than the black crappie (e.g., Carlander 1977; Trautman 1981; Ellison 1984; Etnier and Starnes 1993; Miranda and Lucas 2004). Even though the difference in turbidity tolerance is frequently noted, both crappie species occur in turbid and clear water habitats, and an obvious mechanism or adaptation explaining the apparent difference in tolerance is lacking (e.g., Barefield and Ziebell 1986). Some indirect evidence (e.g., growth, survival) suggests that white crappies can feed more efficiently in turbid waters than black crappies or that white crappies compete poorly in clear waters with other centrarchids (e.g., Carlander 1977; Ellison 1984; Pope 1996). White crappies move extensively, often show distinct diel activity patterns, and can show persistent occupation of home activity areas in the summer. In rivers in Missouri, tagged individuals covered 34 to 42 km in 21 to 91 days (Funk 1957) and others have noted movements up to 30 km (review in Hansen 1951; Siefert 1969a). Increased movement in spring and early summer is attributed to aggregation in spawning areas and postspawning foraging (Guy et al . 1994). Adult white crappies show high levels of nocturnal activity (see section on food), but overall patterns of movement and activity vary seasonally and daily among seasons (e.g., Hansen 1951; Morgan 1954; Greene and Murphy 1974; Markham et al . 1991; Guy et al . 1994). In an Ohio reservoir, diel movement of large white crappie (271–352 mm TL) in summer rapidly increased at dusk when light intensity was zero, peaked at night (average 47 m/h), and declined at dawn. Movement was low throughout the day (average 17 m/h). During the day, the species was associated with steeply sloped bottoms and the presence of structure (e.g., tree stumps, logs, rocks). Individuals tended to occupy deeper water during the day than at night (e.g., 5.4 vs 4.3 m, respectively), generally staying within 0.5 m of the bottom. Median summer home activity areas were 0.49 to 0.63 ha during the day and 1.25 ha at night (Markham et al . 1991). In a shallow, homogeneous glacial lake in South Dakota, movement patterns of large radio-tagged white crappie tracked from April to September were more extensive and less patterned. Over the tracking period, median movement was 73.2 m/h (range: 0–1,523 m/h) and was highest in May (102.1 m/h) and July (82.4 m/h). Diel movement patterns were indistinct or variable, but tended to peak at dawn and dusk. Median home activity area was large relative to the reservoir study (15.8 ha) and varied considerably (range: 0.1–85.0 ha) (Guy et al . 1994). The larger home range, relative to the other study, was attributed to greater foraging demands or the lack of cover and bottom structure in the homogeneous habitat of the lake. Cover or structure tends to hold individuals within a limited area for prolonged periods (Markham et al . 1991; Guy et al . 1994). Food: The white crappie is primarily a midwater, particulate-feeding zooplanktivore and invertivore that shifts to piscivory at a relatively large size (∼ 160 mm TL) compared to other piscivorous centrarchids (O’Brien et al . 1984). Numerous, long gill rakers likely play an important functional role in the extended period of zooplanktivory (Wright et al . 1983). Food of large individuals (>160 mm TL) consists primarily of small fishes (e.g., clupeids, other white crappies and sunfishes, minnows, silversides), zooplankton, immature aquatic insects (e.g., chironomid larvae and pupae, burrowing mayflies), and amphipods (e.g., Hansen 1951; Morgan 1954; Hoopes 1960; Whiteside 1964; Siefert 1969a; Mathur 1972; Greene and Murphy 1974; Ellison 1984; Muoneke et al . 1992). Large white crappies are among the best documented of any centrarchid for their nocturnal feeding and high levels of nocturnal activity (see section on habitat). Large individuals feed at dusk, sporadically throughout the night, and intensively at dawn, feeding very little or not at all during the day (Childers and Shoemaker 1953; Greene and Murphy 1974). In lentic waters, intermediate-size fish (80–150 mm TL) are pelagic zooplanktivores that begin feeding at or near dawn and continue feeding throughout the day (O’Brien et al . 1984; Wright and O’Brien 1984). These pelagic-dwelling individuals can make diel vertical migrations to exploit vertically migrating zooplankton and dipteran larvae and pupae and to respond to changing levels of temperature, light, and DO (O’Brien et al . 1984). Empirical associations of white crappie abundance and abundance of other fishes in wild populations and mesocosm experiments indicate that 130 to 199 mm TL white crappie are highly effective predators that rapidly find and eat larval fishes (e.g., bluegills, walleye). Predation by white crappies is so effective it could drastically limit recruitment of the prey fish species (Kim and DeVries 2001; Quist et al . 2003). Young-of-the-year white crappies feed most heavily during daylight hours on crustacean zooplankton (e.g., copepods and cladocerans) and small dipteran larvae and pupae, but some feeding occurs continuously over a 24-hour period (Siefert 1968, 1969a; Mathur and Robbins 1971; Overmann et al . 1980; DeVries et al . 1998). Individuals can actively search for, pursue, and capture zooplankton prey down to water temperatures of at least 7◦ C (O’Brien et al . 1986).
Centrarchid identification and natural history
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The white crappie is adapted behaviorally and visually for detecting zooplankton prey, but foraging success is affected by prey size, prey movement, light intensity, and turbidity. White crappies use a stereotyped saltatory (pause-travel) search strategy in which they visually locate and attack individual prey. In this strategy, they search briefly for a prey item while stationary and, if they do not locate prey, swim a short distance before stopping to scan again (O’Brien 1979; O’Brien et al . 1986, 1989; Browman and O’Brien 1992). The white crappie retina has a high density of cones in the far temporal region along the eye’s horizontal meridian, an apparent adaptation for detecting open-water zooplankton. Highest probabilities and maximum distances that white crappie will pursue small zooplankters (1–2 mm) are concentrated in a 60-degree forward-directed pie-shaped wedge of limited height (Browman et al . 1990) in which the species is better able to discriminate the absolute size of prey (O’Brien et al . 1985). The wedge-shaped field of maximum foraging corresponds well with the position of the high-density photoreceptor region on the retina (Browman et al . 1990). Under well-lit, lowturbidity conditions (80 lux, 1 NTU), the distance at which individuals (∼ 160 mm TL) can detect prey (reactive distance) increases from about 4 to 30 cm as prey size increases from 1 to 3 mm, and reactive distance for moving prey increases about threefold. For 3-mm prey, white crappie reactive distance is little affected by decreases in illumination from 106 to 10 lux, but from 10 lux to 0.97 lux, reactive distance decreases from about 25 to 6 cm. Differences in reactive distance across prey sizes (1–3 mm) at the lowest light intensities are minimal. Reactive distance to a 2.4-mm prey at 80 lux decreases as an approximate log function of turbidity from about 20 cm at 1 NTU to 5 cm at 33 NTU (Wright and O’Brien 1984). Reproduction: Maturity is usually reached at age 2+ to age 3+ and a minimum size of about 140–180 mm TL, although stunted individuals in dense populations reportedly spawn at 110 mm TL (Morgan 1951a, 1954; Whiteside 1964; Hansen 1951; Siefert 1969a; Trautman 1981). The white crappie is among the earliest, lowest-temperature spawners in the family. The testes and ovaries enlarge and continue developing in the fall and over winter (Morgan 1951b; Whiteside 1964), which is likely an adaptation for early spawning. Spawning occurs at water temperatures of 11 to 27◦ C with most spawning taking place at 16 to 20◦ C. The duration of the spawning period is variable, lasting from 17 to 53 days, and depending on latitude, spawning activity occurs from late March to June or mid-July (Hansen 1951; Morgan 1954; Whiteside 1964; Siefert 1969a; Carlander 1977; McDonough and Buchanan 1991; Pope and DeVries 1994; Travnichek et al . 1996; Sammons et al . 2001). Year-to-year fidelity to nesting areas is not apparent (Hansen 1965). Male white crappies have less fastidious nest-building habits than some centrarchids. Males establish individual territories but apparently do not use caudal sweeping to clear the nesting area. The male remains upright with the abdomen touching or nearly touching the substrate and uses vigorous 3- to 5-second bursts of fin and body movements to sweep out a roughly circular area (about 15–30 cm diameter), actions which remove only the loosest bottom material. Nest-clearing stops before the well-defined depression typical of most centrarchids is created (Hansen 1965; Siefert 1968). Interestingly and atypical among centrarchids, the female often engages in similar nest cleaning behaviors just before spawning and after egg deposition. Substrate at the nest site appears less important to the male than being near some protective cover or bottom vegetation (Siefert 1968). Nests are located on sod clumps, clay, gravel, rock piles, hollows made among aquatic plants, filamentous algae, or roots as well as the surfaces of boulders, rootwads, and submerged brush or trees (Hansen 1943, 1951, 1965; Breder and Rosen 1966). Nests are placed at water depths of 0.1 to 1.5 m (anecdotally up to 6 m, Hansen 1965). Nest spacings suggest colonies (35–50 nests/colony, 46–76 cm apart), and solitary nests are rare (3 of 150), but nests along shorelines (3–15 nests) are in linear arrangements up to 1.2 m apart (Hansen 1965). Nest-guarding males repeatedly repulse approaching females until the female finally stops retreating from the male’s territory when chased, and the male accepts the female (Siefert 1968). The female circles the nest alone but ultimately moves over the bottom of the nest in a head-to-head, broadside position with the male. As both quiver and move forward with vents touching, she slides under the male, causing the pair to move in a curve as gametes are released. Each quivering act lasts about 4 seconds with intervals of 30 seconds to 20 minutes, at which time females often leave the nest. Spawning with a single female can continue from 45 minutes to 2.5 hours (Siefert 1968). In spawning pens, one female spawned in the nest of two different males, and on two occasions an intruding male joined a spawning female and guardian male to steal fertilizations (Siefert 1968). Eggs in two distinct stages of development in two nests suggested that multiple spawnings occurred over a 2-day period (Siefert 1968). Male white crappie remain relatively motionless over the nest and apparently do not engage in rim circling, but do display (opercle flare) to neighboring males or rush and attack (butt, snap, bite) territorially intruding males and females (Hansen 1965; Siefert 1968). During incubation, the male fans the eggs with constant motion of the pectoral fins (Hansen 1943; Breder and Rosen 1966). Fertilized eggs, which are almost completely covered with minute debris, often occur in clumps of three or more and are attached to gravel, leaves, twigs,
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Centrarchid fishes
grass, algae, or plants in and well outside the periphery of and even above the nest (Hansen 1943, 1965; Siefert 1968). Mature ovarian eggs are small, ranging from 0.82 to 0.92 mm in diameter, and fertilized water-hardened eggs average 0.89 mm diameter (Hansen 1943; Morgan 1954; Whiteside 1964). Size-adjusted batch fecundities are higher than any other centrarchid except the black crappie (see accounts on Archoplites and Centrarchus), but female fecundity shows high interannual variation within populations and high variation among populations (Mathur et al . 1979; Dubuc and DeVries 2002; Bunnell et al . 2005). Some females retain ripe eggs throughout the spawning period (Morgan 1954; Whiteside 1964), and gonadosomatic values and larval densities may each show two or more temporally separate peaks (Dubuc and DeVries 2002), patterns which are suggestive of partial release of a single batch over a protracted period, production of two or more batches by a female, or asynchrony in maturation of females. Fecundity increases with female size. The relationship between number of mature eggs (Y) and TL (X) is described by the function log Y = −5.301 + 4.24 log X (formula from data in Morgan 1954, average of 20 length classes, 159–330 mm TL, for 50 females, R2 = 0.87, see also Mathur et al . 1979). At a mean size of 230 cm TL, a female potentially can produce 51,609 mature eggs in a single batch (range: 10,787 eggs at 159 cm TL to 238,506 eggs at 330 cm TL). Hatching occurs in 1.8 to 2.1 days at 18.3 to 19.4◦ C (3.9 days at 14.4◦ C, about 1 day at 22.8◦ C) (Morgan 1954; Siefert 1968). Hatchlings are of 1.22 to 2.74 mm TL, and swim-up larvae disperse on average at 4 days post hatch (range: 2.1 to 6.8 days) at a size of 4.1 to 4.6 mm TL (Morgan 1954; Siefert 1968, 1969b; Sweatman and Kohler 1991; Browman and O’Brien 1992). Male parental care from egg deposition to dispersal typically lasts for 6 days, but, on the basis of developmental information, could range from 4 days at 22 to 23◦ C to 11 days at 14 to 15◦ C (Siefert 1968). Larvae disperse from nesting areas to forage in open water (Siefert 1969a; Overmann et al . 1980). Nest associates: None known. Freshwater mussel host: Confirmed host to A. ligamentina, A. plicata, A. suborbiculata, E. complanata, L. cardium, L. siliquoidea, L. complanata, and L. recta (Young 1911; Lefevre and Curtis 1912; Howard 1914; Coker et al . 1921; Barnhart and Roberts 1997). Putative host to L. reeveiana (unpublished sources in OSUDM 2006). Conservation status: The white crappie is secure throughout its native range (Warren et al . 2000; NatureServe 2006). Similar species: The black crappie has a shorter predorsal region, usually 7 to 8 dorsal spines, and no dark bars on sides. These phenotypic characters are not entirely reliable in separating the two crappie species where both species and their hybrids co-occur (Dunham et al . 1994; Smith et al . 1995). Systematic notes: Pomoxis annularis forms a sister pair with P. nigromaculatus. The pair is basal to a clade comprised of the genera Archoplites and Ambloplites (Roe et al . 2002; Near et al . 2004, 2005). Comparative studies of variation across the range of P. annularis are lacking. Importance to humans: White crappies are a popular sport fish and like black crappies can maintain recruitment and growth that can sustain extremely high levels of exploitation as sport fisheries (e.g., 60% for age 3 and older fish, Colvin 1991). In southern reservoirs, many thousands of crappies are harvested by anglers in the weeks before spawning when fishes, loosely aggregated near cover, go on a feeding spree, perhaps in response to rising water temperatures or preparatory to spawning (Etnier and Starnes 1993; Allen and Miranda 1996; Miranda and Dorr 2000; Dorr et al . 2002). During this time, white crappies are taken easily by anglers using small jigs, streamers, or minnows fished near underwater structure, where fishes are often caught one after the other. Later in spring, white crappies appear most vulnerable to night fishing with minnows below lanterns (Etnier and Starnes 1993).
13.10.2 Pomoxis nigromaculatus (Lesueur) 13.10.2.1 Black crappie Characteristics: See generic account for general characteristics. Deep, extremely compressed body, depth usually 0.37 to 0.45 of SL. Long predorsal region with sharp dip over eye in dorsal profile. Dorsal fin base about equal to or greater than distance from posterior rim of eye to dorsal fin origin. Large, supraterminal, strongly oblique mouth, lower jaw projecting, supramaxilla moderate (≤2 times length of maxilla), upper jaw reaching to or slightly beyond middle of eye. Opercular spot black. Silvery sides profusely speckled and mottled. Long dorsal fin, usually (6)7 to 8(10) spines, 14 to 16 rays; and
Centrarchid identification and natural history
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long anal fin, 6 to 7(8) spines, 16 to 19 rays. Pectoral rays, (13)14(15); vertebrae, 31 to 33(14 + 18 or 19) (Bailey 1938; Keast and Webb 1966; Trautman 1981; Becker 1983; Page and Burr 1991; Etnier and Starnes 1993; Mabee 1993; Jenkins and Burkhead 1994; Smith et al . 1995). Size and age: Typically reach 122 to 160 mm TL at age 1 but first-year growth is highly variable among habitats and apparently less so among latitudes (range, 48–301 mm TL, Carlander 1977). Large individuals measure 300 to 400 mm TL, weigh 400 to 500 g, and reach age 6+ to 8+ (maximum 560 mm TL, 2.72 kg, age 13+) (Carlander 1977; Page and Burr 1991; Etnier and Starnes 1993). World angling record, 2.05 kg, Nebraska and Virginia (IGFA 2006). Coloration: Gray-green above with upper back and silvery blue sides marked with wavy black lines, dark blotches, and green flecks. Silvery below. Dorsal, anal, and caudal fins with many wavy black bands and pale spots. Males become darker during the breeding season (Page and Burr 1991; Etnier and Starnes 1993; Jenkins and Burkhead 1994). The presence of a black predorsal stripe (colloquially known as the black-nose or black-stripe crappie) in some individuals is the expression of a dominant trait controlled by a single gene (Gomelsky et al . 2005). Native range: The native range presumably includes Atlantic Slope drainages from Virginia to Florida, Gulf Slope drainages west to Texas, and the St. Lawrence River-Great Lakes and Mississippi basins from Quebec to Manitoba and south to the Gulf of Mexico (Page and Burr 1991). The wide introduction and establishment of the black crappie renders accurate determination of the native range difficult (Page and Burr 1991; Fuller et al . 1999). As the introduced black crappie became abundant in some California waters, the only native centrarchid, the Sacramento perch, declined or disappeared (Moyle 2002). Historical shifts in distribution and relative abundance suggest that the black crappie has declined or has been replaced by the white crappie because of increased turbidity of waters (e.g., South Dakota, Carlander 1977; Illinois, Smith 1979; Ohio, Trautman 1981; Wisconsin, Becker 1983). In some reservoirs, the black crappie hybridizes extensively with the white crappie (see account on P. annularis). Habitat: The black crappie inhabits lakes, ponds, sloughs, and backwaters and pools of streams and rivers. The species is most common in lowland habitats, large reservoirs, and navigation pools of large rivers but is rare in upland rivers and streams. The black crappie is usually associated with clear waters, absence of noticeable current, and abundant cover (e.g., aquatic vegetation, submerged timber) (Carlander 1977; Werner et al . 1977; Conrow et al . 1990; Page and Burr 1991; McDonough and Buchanan 1991; Keast and Fox 1992; Etnier and Starnes 1993; Pflieger 1997). The species is apparently moderately tolerant of oligohaline conditions, occasionally entering tidal waters (usually