Hormones and Reproduction of Vertebrates - Vol 1: Fishes

Hormones and Reproduction of Vertebrates Hormones and Reproduction Hormones and Reproduction Hormones and Reproductio...

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Hormones and Reproduction of Vertebrates

Hormones and Reproduction Hormones and Reproduction Hormones and Reproduction Hormones and Reproduction Hormones and Reproduction

of Vertebrates, of Vertebrates, of Vertebrates, of Vertebrates, of Vertebrates,

Volume 1dFishes Volume 2dAmphibians Volume 3dReptiles Volume 4dBirds Volume 5dMammals

Hormones and Reproduction of Vertebrates Volume 1: Fishes

David O. Norris Department of Integrative Physiology University of Colorado Boulder, Colorado

Kristin H. Lopez Department of Integrative Physiology University of Colorado Boulder, Colorado

AMSTERDAM BOSTON HEIDELBERG LONDON NEW YORK OXFORD PARIS SAN DIEGO SAN FRANCISCO SINGAPORE SYDNEY TOKYO Academic Press is an imprint of Elsevier

Academic Press is an imprint of Elsevier 32 Jamestown Road, London NW1 7BY, UK 30 Corporate Drive, Suite 400, Burlington, MA 01803, USA 525 B Street, Suite 1800, San Diego, CA 92101-4495, USA First edition 2011 Copyright Ó 2011 Elsevier Inc. All rights reserved Cover images Front cover image: Amphiprion percula, the orange clownfish. Courtesy of iStockphoto: Image 6571184. Back cover image: Atlantic hagfish (Myxine glutinosa) eggs. Courtesy of Stacia A. Sower, University of New Hampshire, Durham, NH, USA and Scott I. Kavanaugh, University of Colorado, Boulder, CO, USA. 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 without the prior written permission of the publisher Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone (+44) (0) 1865 843830; fax (+44) (0) 1865 853333; email: [email protected]. Alternatively, visit the Science and Technology Books website at www.elsevierdirect.com/rights for further information. Notice No responsibility is assumed by the publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. Because of rapid advances in the medical sciences, in particular, independent verification of diagnoses and drug dosages should be made. British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library. Library of Congress Cataloging-in-Publication Data A catalog record for this book is available from the Library of Congress. ISBN: ISBN: ISBN: ISBN: ISBN: ISBN:

978-0-12-374932-1 978-0-12-375009-9 978-0-12-374931-4 978-0-12-374930-7 978-0-12-374929-1 978-0-12-374928-4

(Set) (Volume (Volume (Volume (Volume (Volume

1) 2) 3) 4) 5)

For information on all Academic Press publications visit our website at elsevierdirect.com Typeset by TNQ Books and Journals Pvt Ltd. www.tnq.co.in Printed and bound in United States of America 10 11 12 13 14 15 10 9 8 7 6 5 4 3 2 1

Dedication

Richard Evan Jones

This series of five volumes on the hormones and reproduction of vertebrates is appropriately dedicated to our friend and colleague of many years, Professor Emeritus Richard Evan Jones, who inspired us to undertake this project. Dick spent his professional life as a truly comparative reproductive endocrinologist who published many papers on hormones and reproduction in fishes, amphibians, reptiles, birds, and mammals. Additionally, he published a number of important books including The Ovary (Jones, 1975, Plenum Press), Hormones and Reproduction in Fishes, Amphibians, and Reptiles (Norris and Jones, 1987, Plenum Press), and a textbook, Human Reproductive Biology (Jones & Lopez, 3rd edition 2006, Academic Press). Throughout his productive career he consistently stressed the importance of an evolutionary perspective to understanding reproduction and reproductive endocrinology. His enthusiasm for these subjects inspired all with whom he interacted, especially the many graduate students he fostered, including a number of those who have contributed to these volumes.

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Contents

Series Preface Volume Preface Contributors

xiii xv xvii

1. Sex Determination in Fishes

1

1. Introduction 2. Sex Determination 3. Sexual Differentiation 3.1. Testicular Differentiation 3.2. Ovarian Differentiation 4. Environmental Effects on Sex Determination and Differentiation 5. Conclusions and Future Directions Acknowledgements References

1 2 4 4 7

2. Conserved and Divergent Features of Reproductive Neuroendocrinology in Teleost Fishes 1. Introduction 2. The Unique HypothalamicePituitaryeGonadal (HPG) Axis of Teleosts 3. Gonadotropin-Releasing Hormone (GnRH) 3.1. Discovery of Gonadotropin-releasing Hormone (GnRH) in Teleosts 3.2. Two or Three Gonadotropin-releasing Hormone (GnRH) Genes in Fishes 3.3. Gonadotropin-releasing Hormone (GnRH) Distribution in the Brain of Fishes 3.4. Gonadotropin-releasing Hormone (GnRH)’s Functions are Mediated by Multiple Receptors 3.5. Pulsatile Release of Gonadotropin-releasing Hormone (GnRH) in Fishes? 4. Other Brain Factors Stimulating GTH Release 4.1. Neuropeptide Y (NPY) 4.2. g-aminobutyric Acid (GABA) 4.3. Gonadotropin-inhibiting Hormone (GnIH)

9 10 11 12

15 15 16 17 18 19 19

21 21 22 22 22 23

5. Dopamine (DA), a Brain Inhibitor of Reproduction 5.1. Dual Brain Control of Pituitary Gonadotropins (GTHs) in Teleosts 5.2. Diversity of Dopamine (DA) Roles in the Control of Teleost Reproduction 5.3. Evolutionary Origin of Dopamine (DA) Inhibitory Control of Reproduction in Teleosts 5.4. Modulation of Dopamine (DA) Inhibitory System by Internal Factors 5.5. Modulation of Dopamine (DA) Inhibitory System by Environmental Cues 6. Kiss, a New Actor in the Brain’s Control of Reproduction 6.1. Discovery of the Indispensable Role of Kiss in the Control of Reproduction in Mammals 6.2. Investigation of the Kisspeptin (Kp) System in Teleosts 6.3. Origin and Evolution of the Kisspeptin (Kp) System 6.4. The Kisspeptin (Kp) System in Fishes: Missing Link Between Growth/ Metabolism and Reproduction 7. Sex Steroids in the Brain of Fishes 7.1. Aromatase Expression and High Sexual Plasticity of the Brain in Fishes 7.2. Classical Positive and Negative Feedbacks 7.3. Expression of Other Steroidogenic Enzymes in the Brain of Fishes 8. Conclusions and Perspectives References

3. Testicular Function and Hormonal Regulation in Fishes 1. Introduction 2. Testis Structure and Spermatogenesis: an Overview 3. Testicular Hormones 3.1. Steroids

23 23 24

24 26 26 27

27 27 28

30 31 31 32 34 34 35

43 43 44 47 47

vii

viii

3.2. Additional Hormones and Signaling Molecules 3.3. Genomic Approaches 4. Endocrine Regulation of Testis Structure and Function 5. Temporal Aspects of Testicular Function 5.1. Development from Undifferentiated Gonad to Functional Testis 5.2. Release of Sperm 5.3. Seasonal Aspects of Testicular Function 6. Accessory Gonadal Structures 6.1. Testicular Glands and Testicular Blind Pouches 6.2. Seminal Vesicles 6.2.1. Seasonal variation 6.2.2. Steroidogenesis 6.2.3. Hormonal regulation 6.2.4. Components of seminal vesicle plasma and their functions 6.2.4.1. Ionic concentration, pH, and osmolality 6.2.4.2. Lipids, carbohydrates, and proteins 6.2.4.3. Steroid glucuronides and olfaction 7. Intraspecific Variation in Sperm Characteristics and Testicular Function 8. Conclusions Acknowledgments References

4. Regulation of Ovarian Development and Function in Teleosts 1. Introduction: Fish Models of Reproductive Strategies 2. Morphological Aspects of the Teleost Ovary and Stages of Oocyte Development 3. Differentiation of Primordial Germ Cells into Oogonia 4. Oogenesis, Oocyte Growth, and Development 5. Oocyte Maturation and Ovulation 6. Final Considerations References

5. Thyroid Hormones and Reproduction in Fishes 1. Introduction 2. Thyroid Hormone Delivery 2.1. Regulation of Circulating Thyroid Hormone (TH) Levels

Contents

48 48 49 51 51 51 51 52 53 53 53 53 53 54 54 54 55 55 57 57 58

65 65 66 68 70 73 76 77

83 83 84 84

2.2. Thyroid Hormone (TH) Transporters 2.3. Thyroid Hormone (TH) Clearance 2.4. Thyroid Hormone (TH) Receptors 2.4.1. Nuclear triiodothyronine (T3) receptors 2.4.2. Plasma membrane thyroxine (T4) receptor 3. The Thyroid Tissue of Fishes 3.1. Thyroid Hormone (TH) Synthesis and Release 4. Thyroid Hormone (TH) and Reproduction In Fishes 4.1. Correlative Studies Examining Changes in Thyroid Hormone (TH) Function During Reproductive Maturation 4.2. Identification of Thyroid Hormone (TH) Regulatory Elements in Gonadal Tissue 4.3. Assessment of Thyroid Function Using Sex Steroid Treatment 4.4. Manipulation of Thyroid Function to Induce Changes in the Reproductive System 5. Conclusions References

6. Stress and Reproduction 1. Introduction 1.1. Effectors of the Stress Response 1.2. Effectors of Reproductive Functions 2. Effects of Stress on the Hypothalamice PituitaryeGonadal (HPG) Axis 2.1. Effects of Stress on the Central Nervous System (CNS) 2.2. Effects of Stress at the Level of the Pituitary 2.3. Effects of Stress on Hepatic Vitellogenesis 2.4. Effects of Stress on Gonadal Function 3. Life Stage-Specific Effects of Stress on Reproduction 3.1. Impact of Stress During Embryonic and Larval Stages 3.2. Impacts of Stress on Puberty 3.3. Impacts of Stress on Adults 4. Effects of Sex and Reproduction on the HypothalamicePituitaryeInterrenal (HPI) Axis 5. Reproduction and Resistance to Stress 6. Conclusions References

87 87 87 88 88 89 90 90

91

92 93

94 95 96

103 103 103 104 105 105 106 106 107 108 108 109 109

110 111 112 113

ix

Contents

7. Hormones and Sexual Behavior of Teleost Fishes 1. Theoretical Constructs: Appetitive and Consummatory Phases 2. Patterns of Sexual Behavior 2.1. Defending and Preparing a Spawning Site 2.2. Finding and Choosing a Mate: Species Identification, Sexual Discrimination, and Mate Choice 2.3. Internal and External Fertilization 2.4. Sex-role Reversal 2.5. Alternative Reproductive Tactics 3. Endocrine Mechanisms Regulating Sexual Behavior 3.1. Gonadal Steroids 3.1.1. Gonadal steroids and male sexual behavior 3.1.2. Gonadal steroids and female sexual behavior 3.2. Neuropeptides 3.2.1. Gonadotropin-releasing hormone (GnRH) 3.2.2. Arginine-vasotocin (AVT) 3.2.3. Isotocin (IST) 3.3. Prostaglandins (PGs) 4. Brain Circuits Underlying Sexual Behavior in Fishes 5. Prospects for Future Research 5.1. In-vivo Imaging of Brain Activity 5.2. Molecular Biology 5.3. Genetics Acknowledgements References

8. Neuroendocrine Regulation in Sex-changing Fishes

119 119 120 120

122 124 125 126 127 127 128 131 132 132 133 134 135 135 137 137 137 138 138 139

149

1. Introduction 149 2. Hermaphroditism in Fishes 149 2.1 Protogynous Sex Reversal 151 2.2 Protandrous Sex Reversal 151 2.3 Bidirectional (Serial) Sex Reversal 152 3. Hypotheses of Natural Sex Reversal 152 4. Social Factors Affecting Sex Reversal 153 5. Neuroendocrine Factors Affecting Sex Reversal 154 5.1 Gonadal Steroids 154 5.2 Peptides 156 5.3 Monoamines 157 6. Studies on the Saddleback Wrasse 158 7. Future Research 164 References 164

9. Hormonally Derived Sex Pheromones in Fishes

169

1. Introduction 169 2. Hormonal Pheromones in Fishes 170 2.1. Goldfish and Related Cypriniforms 172 2.1.1. Female preovulatory steroid pheromone 173 2.1.2. Female postovulatory prostaglandin pheromone 174 2.1.3. The male goldfish pheromone(s) 175 2.1.4. Hormonal pheromones in other cypriniforms 176 2.2. Order Salmoniformes 177 2.2.1. Genus Salmo: Atlantic salmon and brown trout 177 2.2.2. Genus Salvelinus (chars) 178 2.2.3. Genus Oncorhynchus (Pacific salmon) 178 2.2.4. Hormonal pheromones and salmonid phylogeny 179 2.3. Order Perciformes 179 2.3.1. The Eurasian ruffe (Gymnocephalus cernuus) 179 2.3.2. The black and round gobies 180 2.3.3. Family Cichlidae 182 3. Hormonal Pheromones and the Issue of Species Specificity 184 Acknowledgements 188 References 188

10. Reproduction in Agnathan Fishes: Lampreys and Hagfishes 1. Introduction 2. Hagfish Reproduction 2.1. The Hagfish Gonad 2.1.1. Sex differentiation 2.1.2. The ovary 2.1.3. The testis 2.2. Secondary Sexual Characteristics 2.3. The Hypothalamicepituitaryegonadal (HPG) Axis of Hagfishes 2.3.1. Neurohypophysis and adenohypophysis 2.3.2. Reproductive hormones 3. Lamprey Reproduction 3.1. The Lamprey Gonad 3.1.1. Lifecycle 3.1.2. Development of larval gonads 3.1.3. The ovary 3.1.4. The testis

193 193 194 195 195 195 196 196 196 196 196 197 197 197 197 198 198

x

3.2. Secondary Sexual Characteristics 3.3. The Hypothalamicepituitaryegonadal (HPG) Axis of Lampreys 3.3.1. Neurohypophysis and adenohypophysis 3.3.2. Gonadotropin-releasing hormone (GnRH) 3.3.3. Gonadotropin-releasing hormone receptor (GnRH-R) 3.3.4. Gonadotropin (GTH) 3.3.5. Glycoprotein hormone (GpH) receptors 3.3.6. Reproductive steroids 3.4. Circulating Hormones During Reproductive Cycles 4. Summary References

11. Hormones and Reproduction in Chondrichthyan Fishes 1. Reproduction in Chondrichthyan Fishes 1.1. Reproductive Modes 1.2. Reproductive Cycles 1.3. Mating and Reproductive Behaviors 2. The HypothalamicePituitaryeGonadal (HPG) Axis 2.1. Gonadotropin-releasing Hormone (GnRH) 2.2. Pituitary Structure and Gonadotropins (GTHs) 3. Hormonal Regulation in Females 3.1. Structure and Function of the Female Reproductive Tract 3.2. Steroidogenesis, Steroidogenic Enzymes, and Steroid Receptors 3.3. Gonadal Steroid Cycling and Functions 3.3.1. 17b-estradiol (E2) 3.3.2. Progesterone (P4) 3.3.3. Androgens 4. Hormonal Regulation in Males 4.1. Testicular Structure and Spermatogenesis 4.2. Steroidogenesis, Steroidogenic Enzymes, and Steroid Receptors 4.3. Gonadal Steroid Cycling and Functions 4.3.1. Androgens 4.3.2. Estrogens 4.3.3. Progesterone (P4) 5. Other Hormones Involved in Reproduction in Males and Females 5.1. Corticosterone (CORT) 5.2. Relaxin

Contents

200 202

5.3. Thyroid Hormones 5.4. Calcitonin (CT) 5.5. Serotonin (5-HT) 5.6. Neurohypophysial Hormones 6. Hormones, Sexual Differentiation, and Sexual Maturation 7. Hormones, Reproductive Behaviors, and Sensory Function 8. Environmental Influences on Circulating Hormone Levels and Reproduction 9. Conclusions and Future Directions Acknowledgements References

203 204 205

12. Hormones and Reproduction of Sarcopterygian Fishes

198 198 198 198 199 199

209 209 209 210 211 211 211 213 215 215 216 217 217 219 219 219 219 220 221 221 222 223 223 223 223

1. Introduction 2. Coelacanths 3. Lungfishes 3.1. Spawning 3.2. Parental Care of Young 3.3. Hormones Involved in Reproduction 4. Concluding Remarks References

224 224 224 225 226 229 229 230 231 231

239 239 240 240 241 241 241 243 243

13. Endocrine-active Chemicals (EACs) in Fishes 245 1. Introduction 1.1. Endocrine Disruption in Fishes 2. Mechanisms of Endocrine-Active Chemical (EAC) Signaling 2.1. Estrogenic Signaling Pathways 2.1.1. Estrogen receptors (ERs) 2.1.2. Estrogen receptor (ER) ligand promiscuity 2.1.3. Estrogen additivity 2.2. Androgenic Signaling Pathways 2.3. The Effect of Endocrine-active Chemicals (EACs) on Steroidogenesis and Steroid Metabolism 3. Multidimensional Mixture Complexity 4. Consequences of Specific Life-Stage Exposures 5. Organizational and Activational Effects of Endocrine-Active Chemicals (EACs) 5.1. Organizational Disruption 5.1.1. Gonadal differentiation 5.1.2. Gonadal intersex 5.2. Activational Disruption

245 246 246 247 247 247 247 248

248 248 248 248 249 249 249 251

xi

Contents

5.2.1. Vitellogenin (Vtg) induction in male fishes 5.2.2. Central activational effects: feedback disruption 5.2.3. Activation of gametogenesis 6. Evidence of Reproductive Disruption in Free-Living Fishes 6.1. Waste Water Treatment Plant (WWTP) Effluent Interaction with Estrogen Signaling Networks 6.1.1. Steroidal estrogens in waste water treatment plant (WWTP) effluents 6.1.2. Synthetic steroidal estrogens in waste water treatment plant (WWTP) effluents

251 252 252 252

253

253

253

6.1.3. Nonsteroidal estrogenic compounds in waste water treatment plant (WWTP) effluent 6.2. Masculinizing Effects of Endocrine-active Chemicals (EACs) 6.3. Pesticides in the Environment 6.4. Neuroactive Pharmaceuticals in the Environment 6.5. Polycholorinated Biphenyls (PCBs) in the Environment 7. Conclusions References Species Index Subject Index Color Plates

253 254 254 255 255 255 256 265 267


Preface

Hormones and Reproduction of Vertebrates Preface to the Series Every aspect of our physiology and behavior is either regulated directly by hormones or modified by their actions, as exemplified by the essential and diverse roles of hormones in reproductive processes. Central to the evolutionary success of all vertebrates are the regulatory chemicals secreted by cells that control sexual determination, sexual differentiation, sexual maturation, reproductive physiology, and reproductive behavior. To understand these processes and their evolution in vertebrates, it is necessary to employ an integrated approach that combines our knowledge of endocrine systems, genetics and evolution, and environmental factors in a comparative manner. In addition to providing insight into the evolution and physiology of vertebrates, the study of comparative vertebrate reproduction has had a considerable impact on the biomedical sciences and has provided a useful array of model systems for investigations that are of fundamental importance to human health. The purpose of this series on the hormones and reproduction of vertebrates is to bring together our current knowledge of comparative reproductive endocrinology in one place as a resource for scientists involved in reproductive endocrinology and for students who are just becoming interested in this field. In this series of five volumes, we have selected authors with broad perspectives on reproductive endocrinology from a dozen countries. These authors are especially knowledgeable in their specific areas of interest and are familiar with both the historical aspects of their fields and the cutting edge of today’s research. We have intentionally included many younger scientists in an effort to bring in fresh viewpoints. Topics in each volume include sex determination, neuroendocrine regulation of the hypothalamuse pituitaryegonadal (HPG) axis, separate discussions of testicular and ovarian functions and control, stress and reproductive function, hormones and reproductive behaviors, and comparisons of reproductive patterns. Emphasis on the use of model species is balanced throughout the series with comparative treatments of reproductive cycles in major taxa.

Chemical pollution and climate change pose serious challenges to the conservation and reproductive health of wildlife populations and humans in the twenty-first century, and these issues must be part of our modern perspective on reproduction. Consequently, we have included chapters that specifically deal with the accumulation of endocrinedisrupting chemicals (EDCs) in the environment at very low concentrations that mimic or block the critical functions of our reproductive hormones. Many authors throughout the series also have provided information connecting reproductive endocrinology to species conservation. The series consists of five volumes, each of which deals with a major traditional grouping of vertebrates: in volume order, fishes, amphibians, reptiles, birds, and mammals. Each volume is organized in a similar manner so that themes can be easily followed across volumes. Terminology and abbreviations have been standardized by the editors to reflect the more common usage by scientists working with this diverse assembly of organisms we identify as vertebrates. Additionally, we have provided indices that allow readers to locate terms of interest, chemicals of interest, and particular species. A glossary of abbreviations used is provided with each chapter. Finally, we must thank the many contributors to this work for their willingness to share their expertise, for their timely and thoughtful submissions, and for their patience with our interventions and requests for revisions. Their chapters cite the work of innumerable reproductive biologists and endocrinologists whose efforts have contributed to this rich and rewarding literature. And, of course, our special thanks go to our editor, Patricia Gonzalez of Academic Press, for her help with keeping us all on track and overseeing the incorporation of these valuable contributions into the work. David O. Norris Kristin H. Lopez

xiii


Preface

Preface to Volume 1 Fishes Fishes represent the most diverse and abundant paraphyletic category of vertebrates. They also have experienced the longest evolutionary history. The endocrine system of fishes is the prototype vertebrate system; it has undergone extensive evolution within the fishes and has provided the basic foundation for the reproductive endocrinology of all vertebrates. Additionally, fishes are of great economic importance for humans as they provide billions of tons of food annually. Unfortunately, in spite of their seeming abundance, the numbers and diversity of fishes are decreasing in the face of persistent human impacts, not only because of their commercial uses but as a result of extensive chemical and thermal pollution of freshwater and marine habitats. Our understanding of fish reproductive endocrinology is basic not only to understanding the reproductive endocrinology of other vertebrates but also to the continued survival of fishes. In this volume, we focus on the basic components of the reproductive endocrinology of fishes, setting the pattern we have followed in all subsequent volumes of this series.

We begin by looking at sex determination, neuroendocrine regulation of the hypothalamicepituitaryegonadal axis, and basic aspects of ovarian and testicular function. Additional chapters deal with the roles of pheromones and stress, and the effects of thyroid function on reproduction of fishes, as well as the endocrine control of reproductive behaviors, including sex reversal, exhibited by certain teleosts. Special chapters deal with hormones during the reproductive cycles of agnathan fishes (lampreys, hagfishes), sarcopterygian fishes (lungfishes), and chondrichthyan fishes (sharks and their relatives). The teleost fishes, which represent the largest segment of living fishes by far, are also the best studied. Consequently, they are thoroughly covered throughout the basic chapters in this volume. The final chapter in this volume addresses the growing problem of disruption of the fish reproductive system by hormone mimics and antagonists, which are appearing in the aquatic environment as a consequence of widespread chemical pollution and hence are capable of reducing the reproductive potential of fishes on a worldwide basis.

xv


Contributors

Nicholas J. Bernier

Masaru Nakamura

University of Guelph, Guelph, ON, Canada

Solution-Oriented Research for Science and Technology (SORST),

Sharon L. Carlisle

Kawaguchi, Saitama, Japan; University of the Ryukyus, Motobu,

University of Oklahoma, Norman, OK, USA

Sylvie Dufour Museúm National d’Histoire Naturelle, Paris, France

Meghan L.M. Fuzzen University of Guelph, Guelph, ON, Canada

James Gelsleichter University of North Florida, Jacksonville, FL, USA

David M. Gonçalves

Okinawa, Japan

David O. Norris University of Colorado at Boulder, Boulder, CO, USA

Rui F. Oliveira Instituto Superior de Psicologia Aplicada, Lisboa, Portugal; Instituto Gulbenkian de Cieˆncia, Oeiras, Portugal

Bindhu Paul-Prasanth Solution-Oriented Research for Science and Technology (SORST), Kawaguchi, Saitama, Japan; National Institute for Basic Biology,

Instituto Superior de Psicologia Aplicada, Lisboa, Portugal, Universidade

Okazaki, Japan

do Algarve, Faro, Portugal

Jason C. Raine

Jean M.P. Joss

University of Saskatchewan, Saskatoon, SK, Canada

Macquarie University, Sydney, NSW, Australia

E. Rocha

Olivier Kah

University of Porto, Porto, Portugal

Universite´ de Rennes, Rennes, France

M.J. Rocha

Hiroshi Kawauchi

University of Porto, Porto, Portugal; Superior Institute of Health

Laboratory of Molecular Endocrinology, Sendai, Miyagi, Japan

Sciences-North (ISCS-N), Paredes, Portugal

Rosemary Knapp

Stacia A. Sower

University of Oklahoma, Norman, OK, USA

University of New Hampshire, Durham, NH, USA

Earl T. Larson

Norm Stacey

Northeastern University, Boston, MA, USA

University of Alberta, Edmonton, AB, Canada

Karen P. Maruska

R. Urbatzka

Stanford University, Stanford, CA, USA

University of Porto, Porto, Portugal

Yoshitaka Nagahama Solution-Oriented Research for Science and Technology (SORST),

Alan Milan Vajda University of Colorado at Denver, Denver, CO, USA

Kawaguchi, Saitama, Japan; National Institute for Basic Biology,

Glen Van Der Kraak

Okazaki, Japan

University of Guelph, Guelph, ON, Canada

xvii


Chapter 1

Sex Determination in Fishes Bindhu Paul-Prasanth,*, y Masaru Nakamura*, ** and Yoshitaka Nagahama*, y *

Solution-Oriented Research for Science and Technology (SORST), Kawaguchi, Saitama, Japan, y National Institute for Basic Biology, Okazaki, Japan, University of the Ryukyus, Motobu, Okinawa, Japan

**

SUMMARY Among the vertebrates, teleost fishes display the greatest diversity of sexual phenotypes, thus providing an excellent model to investigate molecular mechanisms of sex determination/differentiation. Sex in fishes is determined by genetically or environmentally based signals. The second vertebrate sex-determining gene, dmy/dmrt1by, was discovered in medaka (Oryzias latipes) but was found to be present only in two Oryzias species, illustrating the vast diversity of sex-determining genes in fishes. By contrast, molecular mechanisms involved in sexual differentiation appear to be conserved among fishes. Several factors have been identified: Gsdf/Dmrt1 for testicular differentiation and Foxl2/ Cyp19a1a/estrogens for ovarian differentiation. Among these factors, Gsdf is specific to the fish lineage and a new player in the field of sexual differentiation of fishes. These factors may play major roles in both genetic and environmental modes of gonadal sex differentiation.

1. INTRODUCTION Most vertebrates adopt a sexual mode of reproduction. Sexually reproducing organisms generally exist in one of two anatomical and physiological states, namely male or female. Males develop testes, while females develop ovaries for production of gametes: sperm and ova, respectively. Development of testes or ovaries is the outcome of differential morphogenesis of the gonadal primordium. Essentially, differential morphogenesis is established through repression of the manifestation of one sex and promotion of the other sex by sex determination (SD). Vertebrates exhibit a wide array of sex-determining mechanisms, sometimes even at the species level, ranging from environmental sex determination (ESD) to genetic sex determination (GSD). Among vertebrates, fishes are the only group that possesses all the various mechanisms of sex determination known for vertebrates. Therefore, study of sex determination using various fishes as animal models constitutes an essential and exciting part of the field of sex determination. Hormones and Reproduction of Vertebrates, Volume 1dFishes Copyright Ó 2011 Elsevier Inc. All rights reserved.

Sex determination kicks off the program for sexual development in the undifferentiated gonadal primordium and sex differentiation orchestrates further structural development of the gonads according to their designation at the SD step. In general, mechanisms underlying sex differentiation are more or less conserved across genera. During sexual differentiation, the somatic as well as the germ cells in the gonads differentiate and the gonads undergo testis- or ovary-specific morphogenesis. In fishes, these processes are controlled largely by genetic and hormonal factors. However, most of the earlier studies describing the involvement of steroid hormones in sexual differentiation are based on the exogenous administration of sex steroids. Since numerous reviews are available on this subject, we have not dealt much with this topic in the present chapter. Instead, we will discuss the role of endogenous sex steroids as revealed by studies on the expression patterns of steroidogenic enzymes during early sexual differentiation stages. A comprehensive state-of-the-art analysis of the field concerning sex determination and differentiation among fishes was done by Devlin and Nagahama in 2002, the year in which the first male sex-determining gene among fishes was identified from a small teleost fish, the Japanese medaka (Oryzias latipes) (Matsuda et al., 2002; Nanda et al., 2002). This review extensively covered about three decades of research prior to 2002 and gave elaborate descriptions of almost all aspects of fish sex determination/ differentiation; e.g., various modes of sex determination, gonadal differentiation and development, endocrine regulation of gonadal differentiation and morphogenesis, etc. More recently, several prominent studies in the field of sex determination among fishes have unraveled more facts, factors, and mechanisms involved at the molecular, cellular, and physiological levels, filling in the gaps in the evolution of sex-determining mechanisms. Use of modern molecular tools such as transgenesis and gene knock-down have made it possible to study the cellular and molecular processes involved in the early stages of gonadal development among 1

2

fishes so as to reveal the evolutionary relationships among different vertebrate classes. Some of the genes found in fishes have similar roles in higher vertebrates, while some possess reversed roles. Here, we have reviewed mainly more prominent studies that have come out within and after the year 2002. Though fishes include innumerable species, emphasis will be given mostly to model organisms such as medaka, tilapia (Oreochromis niloticus), zebrafish (Danio rerio), and rainbow trout (Oncorhynchus mykiss), as most of the studies on sex determination/differentiation are centered on these species.

2. SEX DETERMINATION Fishes have both XX/XY and ZZ/ZW heterogametic modes of sex determination. In the XX/XY system, males carry the heteromorphic sex chromosomes and the presence of the Y-chromosome triggers gonadogenesis into the testicular pathway. Although a ZZ/ZW system, with females carrying the heteromorphic chromosomes, has been reported in several fishes belonging to the Teleostei (Anguilliformes, etc.), only a limited number of studies dealing with this topic are available. Therefore, for practical reasons, the ZZ/ZW system is not analyzed in detail in this review. In vertebrates with GSD, undifferentiated gonads develop strictly under the control of the programs within their genomes. According to the master regulator positioned at the top of the sex differentiation cascade, gonads develop along one of two alternate pathways, either testis or ovary. In mammals, the SD gene, SRY/Sry (sex-determining region of the Y chromosome), initiates testicular differentiation through the sex-specific upregulation of Sox9, which in turn promotes the testis-specific differentiation of the somatic cell lineages and, thus, the male-specific gonadal architecture (Sekido, Bar, Narvaez, Penny, & LovellBadge, 2004; Kidokoro et al., 2005; Sekido & LovellBadge, 2008). Concomitant to the promotion of the testicular cascade, Sox9 represses the alternate cascade of development by suppressing key genes involved in ovarian differentiation (Wilhelm et al., 2009). Hence, the role of the SD gene is to promote the expression of those factors required for the establishment of the sex-specific gonadal architecture, which will in turn support the differentiation, development, and maturation of the gamete that the gonad produces. The first decade of the twenty-first century has witnessed a major breakthrough in the field of fish sex determination. Matsuda et al. (2002) and Nanda et al. (2002) independently identified the first male sex-determining gene among fishes, dmy (DM-domain gene on the Y chromosome)/dmrt1bY from medaka. It was named so because dmy/dmrt1bY was found to be a duplicate copy of the well-conserved downstream gene, Dmrt1 (doublesex


and mab-3 related transcription factor 1), of vertebrate sexual differentiation on the Y-chromosome. dmrt1 carries the DM domain, which is characterized as the zinc finger-like DNA-binding motif, and the DM domain was originally found in two separate phyla, insects and nematodes: D for doublesex in Drosophila melanogaster and M for male abnormal-3 in Caenorhabditis elegans. These two genes control sexual development in these invertebrates and, more interestingly, DM domain genes have a conserved role in sex differentiation across various phyla. dmy was the second sex-determining gene to be identified among vertebrates, almost a decade after the discovery of the mammalian SRY/Sry (Sinclair et al., 1990; Koopman, Gubbay, Vivian, Goodfellow, & Lovell-Badge, 1991). Though both dmy/dmrt1bY and SRY/Sry are functionally analogous, no structural similarities have been noticed between them. Unlike SRY/Sry, which appears to be unique to mammals, the involvement of DM-domain genes in sex determination appears to be more highly conserved, as evidenced by two recent discoveries. In Xenopus laevis, DM-W has been identified as the sex-determining gene (Yoshimoto et al., 2008). In contrast to dmy, DM-W is involved in ovarian determination. More recently, dmrt1 has been revealed as the male sex-determining gene of chickens (Smith et al., 2009), in which the male carries homomorphic chromosomes, ZZ, and two copies of dmrt1 likely are required for testicular differentiation, unlike in medaka. Thus, the DM domain is involved in the sex determination of animals belonging to three out of five classes that constitute the phylum chordata: fishes, amphibians, and birds. Dmy has been found in only two species of Oryzias: Oryzias latipes (Matsuda et al., 2002; Nanda et al., 2002) and O. curvinotus (Matsuda et al., 2003; Kondo, Nanda, Hornung, Schmid, & Schartl, 2004) and not in any other genus of fishes. Though the medaka follows an XX/XY heterogametic mode of sex determination, the X and Y chromosomes are homomorphic, unlike in mammals. To date, the only difference noted between X and Y chromosomes of O. latipes is the exclusive presence of dmy on the Y chromosome. Accordingly, a natural mutant XY fish in which there was a single insertion in exon 3 of dmy resulting in truncation of its protein showed a female phenotype (Matsuda et al., 2002). Similarly, another XY mutant female showed reduced expression of dmy and a high proportion of XY females were found among the offspring of this mutant female. In addition to these observations, which gave important clues about the sexdetermining role of Dmy, collection of new data in recent years has provided further insights about the functions of Dmy protein. In order to understand the role of Dmy during initiation of testicular differentiation in medaka, a functional analysis

Chapter | 1

3

Sex Determination in Fishes

was carried out by knocking it down with engineered peptide nucleic acid (gripNA) (Paul-Prasanth et al., 2006). dmygripNA and (human cAMP response elementebinding) hCREB-gripNA were injected into one-cell stage medaka embryos. Simultaneously, we treated another batch of the fertilized eggs with 17b-estradiol (E2) to compare the process of female sexual differentiation in these larvae with that of the knock-down larvae, as the mechanisms behind ovarian differentiation could be different. Ovarian differentiation in E2-treated XY larvae was found to be initiated in the presence of an unaltered Dmy (Suzuki, Nakamoto, Kato, & Shibata, 2005); however, the gonads of dmy-gripNA-XY larvae were expected to undergo ovarian differentiation like the normal genetic females as Dmy was not supposed to be present in knock-down embryos. The fry collected on the day of hatching were subjected to both histological (for germ cell counting) and whole-mount in situ hybridization analyses. The sex of the fry was assessed by genomic polymerase chain reaction (PCR) of the DNA extracted from the head or tail region of each fry, using common primers for dmrt1 and dmy. Some of the 0 days after hatching (dah) dmy knock-down XY fry had germ cell numbers comparable to those of the normal XX fry. However, the germ cell numbers in the E2-treated XY gonads at 0 dah were found to be similar to those of the hCREB (control) gripNA-treated and normal XY gonads of the same stage (Figure 1.1). Wholemount in-situ hybridization with the meiosis-specific marker gene, synaptonemal complex protein 3 (scp3), revealed that meiosis was induced in the gonads of the 0 dah dmy knockdown (Figure 1.1) and E2-treated XY fry, but not in the control XY fry. These findings revealed that the mitotic and meiotic activities of the germ cells in the 0 dah dmy knockdown XY larvae were similar to those of the normal XX larvae, suggesting that the microenvironment of these XY gonads is similar to that of the normal XX gonad, where

Dmy is naturally absent. Conversely, E2 treatment failed to initiate mitosis in the XY gonad, possibly due to an active Dmy, even though it could initiate meiosis. This study showed that germ cells in the XY gonad can resume mitotic activity if dmy is knocked down. Corroborating this data, Herpin et al. (2007) found that injection of dmy/dmrt1bY morpholino resulted in increased rate of germ cell proliferation in XY embryos of medaka. These results demonstrate that Dmy is sufficient for male development in medaka, and suggest that the functional difference related to sex differentiation between the X and Y chromosomes in medaka is a single gene. Over-expression of the 117-kb bacterial artificial chromosome (BAC) clone harboring the dmy genomic region in medaka embryos initiated testicular differentiation in the gonads of genetic females (Matsuda et al., 2007). Overexpression of dmy led germ cells in XX gonads into mitotic as well as meiotic arrests. In the dmy over-expression experiments, the total number of germ cells at 0 dah was significantly reduced in both the XX and XY fry. In the 5 dah XX fry injected with dmy, not only was the total germ cell number reduced but the number of germ cells at various stages of development was also reduced. As the total number of germ cells reflects the outcome of active mitosis, the reduced number of germ cells in the transgenic fry may be due to a signal or signals from the surrounding somatic cells that express dmy. The over-expression of dmy using the dmy genomic region induced sex reversal in XX gonads. However, since the genomic region used was large (117-kb), it could not be ruled out that a region within this 117-kb DNA segment but outside the dmy open reading frame (ORF) might be involved in the induction of sex reversal. Therefore, to investigate the ability of the dmyORF to induce sex reversal, we constructed over-expression vectors in which cytomegalovirus (CMV) promoter controlled the dmy or

FIGURE 1.1 Change in germ cell number and meiosisspecific marker gene expression in medaka XY embryos after dmy knock-down. dmy was knocked down using gripNAs in the XY embryos of medaka in order to ascertain its role during early stages of testicular development. dmy and human CREB (control) gripNAs were microinjected into single-cell-stage embryos, and the status of sexual differentiation in the gonads of these embryos was examined on the day of hatching by 80 histology and whole-mount in-situ hybridization XY(WMISH). XY and XY-hCREB fry had fewer germ cells, dmy while XX and XY-dmy possessed more germ cells, indicating initiation of ovarian differentiation in the gonads of the latter groups. Another indication of initiation of ovarian differentiation is the initiation of meiosis exclusively in 0 dah XX gonads. Here, the initiation of meiosis was assessed by WMISH with scp3 (photomi0 crographs). No signals could be detected for scp3 in XYXX XYXYXY hCREB dmy hCREB (upper panel), while strong expression of scp3 was observable in the gonadal region of XY-dmy (lower panel), confirming the entry of germ cells in these gonads into meiosis in the absence of Dmy. See color plate section. 160

Germ cell number

XYhCREB

4

dmrt1 cDNA. These constructs were injected into one-cellstage medaka embryos of the Hd-rR strain, and the injected embryos were reared until the secondary sexual characteristics became apparent. Female-to-male sex-reversed fish were obtained from the embryos that over-expressed dmy, whereas no sex-reversed fish were obtained from the embryos that over-expressed dmrt1. The sex-reversed medaka were fertile. Very recent evidence (Otake et al., 2009) indicated that the exogenous dmy in the above experiments was located on linkage group (LG) 23 in one strain and 5 in another strain, whereas the endogenous dmy was originally located on LG 1. Male development in those fish carrying the heritable artificial chromosomes with exogenous dmy demonstrates that this gene is indeed involved in initiation of testicular differentiation. Localization of the signals for Dmy exclusively in the somatic cells sequestering the primordial germ cells (PGCs) of stage 36 XY embryos suggests that it may regulate further differentiation of PGCs directly or indirectly in these embryos to prevent female sex differentiation (T. Kobayashi, H. Kobayashi, & Nagahama, 2004). In mammals, the role of SRY/Sry is considered to be critical for the upregulation of Sox9 and thus Sertoli cell differentiation (Sekido et al., 2004; Kidokoro et al., 2005; Sekido & Lovell-Badge, 2008). Recent evidence from mice has elucidated the molecular mechanism involved in this process. Sry along with steroidogenic factor-1 (SF-1) bind to multiple elements within the Sox9 gonad-specific enhancer region on its promoter in order to upregulate Sox9 expression in a male-specific manner (Sekido & Lovell-Badge, 2008). Loss- and gain-of-function experiments have shown that Dmy is necessary and sufficient for initiating testicular differentiation in the primordial gonads of genetically male medaka (Paul-Prasanth et al., 2006; Matsuda et al., 2007). However, the molecular mechanisms underlying the action of Dmy remain largely unknown, even after more than half a decade since its discovery, mainly because of lack of knowledge regarding its target genes. Ample evidence collected hitherto indicates that this gene is involved directly or indirectly in the regulation of germ cell proliferation. No other clues have revealed the precise function of Dmy. Other species in which intensive searches for an SD gene are being carried out include tilapia, zebrafish, fugu (Takifugu rubripes), three-spined stickleback (Gasterosteus aculeatus), and rainbow trout; however, to date studies have not pinpointed a single factor as the SD gene in these species, although several candidate genes were identified. Studies have been able to locate only the SD region, but the genes are not yet deciphered (fugu: Kikuchi et al. (2007); rainbow trout: Woram et al. (2003) and Alfaqih, Brunelli, Drew, and Thorgaard (2009); threespined stickleback: Peichel et al. (2004), Ross and Peichel (2008), Ross, Urton, Boland, Shapiro, and Peichel (2009),


Kitano et al. (2009), and Leder et al. (2010)). Thus, the status of dmy as the only SD gene among fishes continues.

3. SEXUAL DIFFERENTIATION Gonadal morphogenesis is initiated soon after the PGCs are translocated to the gonadal primordium. The process of morphogenesis is initiated by species-specific sexdetermining mechanisms. In gonochoristic teleosts such as tilapia, medaka, etc., PGCs start to proliferate (medaka: stage 38 (Kobayashi et al., 2004); tilapia: 8 dah (Kobayashi, Kajiura-Kobayashi, Guan, & Nagahama, 2008)) female-specifically and enter into meiosis, giving a distinct morphology to the gonads of genetic females. On the other hand, genetic males of these species carry gonads at this time that appear undifferentiated at the morphological level as the PGCs remain mitotically and meiotically silent.

3.1. Testicular Differentiation In most fish species, testicular differentiation remains morphologically inconspicuous during initial stages of gonadal development. In the case of medaka, the gonads of genetic males look undifferentiated up to 10 dah, because the germ cells remain mitotically and meiotically quiescent. At 10 dah, the gonads in this species show the first morphologically differentiated testis-specific structure, the acinus, which is the precursor of the seminiferous tubule. This process is comparable to the process of testicular cord formation in mammals. In general, morphologically discernable testicular differentiation is delayed among fishes but there exists much diversity in the developmental stage at which the presumptive testes exhibit the first sign of morphological sex differentiation. However, accumulating data from medaka indicate that the first event during the course of gonadal differentiation in fishes is the differentiation of Sertoli cells. Numerous studies on fishes have shown that treatment of XX larvae with androgens induces sex reversal toward testicular development. However, no direct experimental evidence has been provided to prove whether endogenous androgens have an essential role in initial testicular differentiation in fishes. In tilapia, mRNAs of most steroidogenic enzymes such as cholesterol side-chain cleaving enzyme (P450scc ¼ CYP11A1), 3b-hydroxysteroid dehydrogenase (3b-HSD), 17a-hydroxylase (P45017a ¼ CYP17a1), and 17b-hydroxysteroid dehydrogenase type 1(17b-HSD1) were constantly detected in XY gonads at 5e7 dah (Ijiri et al., 2008). However, unlike in XX gonads, in XY gonads there were no further increases in these levels at 10e25 dah. Another important observation in this study is that the expression of cytochrome P450 11b-hydroxylase

Chapter | 1

5


(P45011b ¼ CYP11B2)dwhich contributes to the synthesis of 11-ketotestosterone (11-KT), the most potent androgen in teleost fishes, from testosterone (T)dis not detected in either XX or XY gonads at 5e25 dah. These findings may indicate that 11-KT is not produced in tilapia gonads until 25 dah and, thus, androgens do not appear to play a major role in testicular differentiation of this species. In contrast, at 70 dah, the mRNA levels of four steroidogenic enzymes including P45011b are higher in XY gonads than in XX gonads (Ijiri et al., 2008). As spermatogenesis is initiated during this period, increased steroidogenic activity of XY gonads may be involved. A recent investigation in rainbow trout confirmed that initiation of testicular differentiation does not require androgen production (Vizziano, Randuineau, Baron, Cauty, & Guiguen, 2007). Further studies are necessary to confirm whether androgen is involved in testicular differentiation in fishes or not. Our recent studies revealed a new factor, gonadale soma-derived factor (Gsdf), associated with testicular differentiation in medaka (Shibata et al., 2010) (Figure 1.2). Originally identified from rainbow trout as a unique member of the transformation growth factor superfamily, Gsdf is a comparatively new player in the field of sexual differentiation of fishes (Sawatari, Shikina, Takeuchi, & Yoshizaki, 2007). Phylogenetic analyses encompassing Gsdfs from several other fish species such as fugu, zebrafish, threespined stickleback, and medaka, along with rainbow trout, have revealed that this gene is specific to the fish lineage. There are two forms of Gsdf in rainbow troutdGsdf1 and -2dwhile only one has been found in medaka (Lareyre et al., 2008). Unlike dmy, expression of the gsdf gene is found in the somatic cells surrounding the germ cells, not only in the XY gonads but also in the XX gonads, especially in granulosa cells around previtellogenic oocytes. However, prior to the onset of early signs of testicular differentiation, a rise is noticed in expression levels of gsdf in XY gonads at

XY

around six days post-fertilization (dpf) (Shibata et al., unpublished). Such a rise is not observed in XX gonads; rather, the expression of gsdf becomes clearly visible in XX gonads only from 10 dah. Colocalization studies have demarcated two types of somatic cells in the XY gonads: one type that expresses only dmy protein and another type showing expression of both dmy and gsdf. The cells expressing both dmy and gsdf could be undergoing differentiation into Sertoli cells, and it is possible that this process precedes all the other morphological changes that have been recorded so far. Therefore, higher levels of gsdf mRNA in somatic cells of XY gonads than that of the XX gonads could be critical for the masculinization of XY somatic cells. Gsdf seems to play a role equally important to Dmy in driving the XY gonads towards testicular differentiation. Expression of dmy is linked to the genotype of the gonads, not to the phenotype. Accordingly, in XY females generated by exogenous E2 treatment, dmy expression has been detected in granulosa cells, suggesting that the presence of dmy is not detrimental to ovarian development (Suzuki et al., 2005). In contrast to dmy, expression of gsdf is closely linked to the gonadal phenotype, because E2 administration downregulated the expression levels of gsdf mRNA in exposed XY embryos (Shibata et al., unpublished). It is possible that Gsdf is the first target of estrogenic chemicals in the male sex-differentiation cascade in medaka. Among fish species, dmrt1 shows a consistent pattern of expression with an exception in medaka. In tilapia (Guan, Kobayashi, & Nagahama, 2002; Ijiri et al., 2008), rainbow trout (Vizziano et al., 2007), and zebrafish (Jørgensen, Morthorst, Anderson, Rasmussen, & Bjerregaard, 2008), expression of dmrt1 coincides with initiation of testicular differentiation (Figure 1.3). Conversely, in medaka, dmrt1 expression becomes testis-specific only at around 20 dah, which is well down the pathway of the initiation of testicular differentiation (Kobayashi et al., 2004). More

XX

FIGURE 1.2 Expression pattern of gsdf mRNA during early stages of sexual differentiation in medaka. Expression pattern of gsdf was examined in the XY and XX gonads on the day of hatching by in-situ hybridization using amplified gsdf aRNA probe. Predominant expression of gsdf was detectable in the somatic cells surrounding the germ cells in the XY gonads. Its expression was almost undetectable in the XX gonads of the same stage. Strong expression of gsdf only in the somatic cells of the presumptive testis during early sexual differentiation stages suggests a role for it in the initiation of Sertoli cell differentiation. See color plate section.

6


dmrt1

ad4bp/sf-1

cyp19a1a

5

foxl2

6

7

5

6

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Days after hatching FIGURE 1.3 Expression patterns of key genes involved in male and female sex differentiation in tilapia. Levels of dmrt1, ad4bp/sf-1, cyp19a1a, and foxl2 mRNAs were quantitatively assessed in monosex populations of tilapia using real-time polymerase chain reaction (PCR). dmrt1 shows a sexually dimorphic expression pattern in tilapia during very early stages of sexual differentiation. No significant difference was noticeable in the expression pattern of dmrt1 at 5 days after hatching (dah), when the gonads remain sexually undifferentiated in this species. However, from 6 dah a dramatic increase in dmrt1 levels was easily discernable only in the genetic males (black circles). On the other hand, ad4bp/sf-1 does not show sexual dimorphism in its expression during early sexual differentiation. Transcripts of cyp19a1a increase between 5e6 dah only in genetically female gonads (open circles), reaffirming the crucial role of 17b-estradiol (E2) in ovarian differentiation in tilapia. A concomitant increase in the expression levels of foxl2 was apparent only in XX gonads at 5e6 dah (open circles).

interestingly, during temperature-induced masculinization of genetic females with no dmy/dmrt1bY in their genomes, dmrt1 expression is upregulated in 50% of the exposed embryos from stage 36, which is much earlier than its usual temporal expression pattern, and, concordantly, 40% of embryos developed testes, suggesting that Dmrt1 in this case acts as the master regulator to initiate testicular differentiation in the absence of Dmy (Hattori et al., 2007). Most recently, in-vitro and in-vivo functional analyses of Dmrt1 in tilapia revealed the possible mechanism by which this gene orchestrates testicular differentiation in genetic males of this species and probably other vertebrates (Wang et al., 2010). Overexpression of dmrt1 driven by CMV promoter in genetically female tilapia embryos

caused retardation of ovarian cavity formation, follicular degeneration, and partial to complete sex reversal. Transcription of the ovarian aromatase gene, P450aro (¼ CYP19A1), was repressed in these embryos, resulting in reduced serum E2 levels. In-vitro analysis revealed that Dmrt1 protein could directly bind to the palindrome sequence, ACATATGT, on the promoter region of P450aro, suggesting further that Dmrt1 probably induces the male pathway through its direct blocking action on estrogen production. Medaka and tilapia have two sox9 genes, sox9a1 and sox9a2. Of these, only sox9a2 is linked with testicular differentiation in these fishes (Nakamoto, Suzuki, Matsuda, Nagahama, & Shibata, 2005; Ijiri et al., 2008). Although sox9a2 is expressed in the somatic cells in the gonads of both XY and XX fishes from stage 33, which is before the onset of Dmy expression, its expression becomes male-specific only from 10 dah. This occurs at around the same stage as acinus formation, leading to the hypothesis that Sox9a2 is involved in the process of acinus formation (Nakamoto et al., 2005; Nakamura et al., 2008). Expression of the sex-determining gene, SRY/Sry, and the upregulation of SOX9/Sox9 in mammals are closely associated with testicular cord formation, while timings of the initiation of expression of the sex-determining gene of medaka, dmy, and acinus formation are not synchronized, leading to the assumption that these two processes are not directly correlated. However, unlike Sry, dmy expression does not decline in XY gonads until at least 10 dah, suggesting that dmy and sox9a2 might be involved with Sertoli cell differentiation and first male-specific structure formation, which are very prolonged processes in medaka. Hence, it is still valid to consider sox9a2 as having a role during early stages of male gonadogenesis in fishes. However, the mechanisms or other cofactors involved remain largely unexplored. In addition to this purported role, sox9a2 is useful in tracing the lineage of somatic cells in both the testis and ovary of medaka (Nakamura et al., 2008). In sox9a2-EGFP (enhanced green fluorescent protein), transgenic medaka testes sox9a2-expressing cells develop as Sertoli cells, while in ovaries the same cells develop as granulosa cells. In other species of fishes such as rainbow trout (Vizziano et al., 2007) and the rice field eel (Monopterus albus) (Lu, Cheng, Guo, & Zhou, 2003; Zhou et al., 2003), sox9a2 is associated with testicular differentiation. Anti-Mu¨llerian hormone (AMH) is a glycoprotein belonging to the transforming growth factor-b superfamily. A major function of AMH in mammals is to mediate regression of Mu¨llerian ducts in males. Targeted mutagenesis has shown that AMH is not required for testicular determination in mice (Behringer, Finegold, & Cate, 1994). Importantly, unlike other vertebrates, fishes do not have Mu¨llerian ducts, indicating a different role for this gene in

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7


fishes. In tilapia, amh mRNA starts to be expressed in indifferent gonads of both sexes, and the expression is upregulated in male gonads but downregulated in female gonads (Ijiri et al., 2008). The sexually dimorphic expression of amh in fish gonads during sex differentiation also has been reported in the Japanese flounder (Paralichthys olivaceus) (Yoshinaga et al., 2004). In these studies, amh expression starts in supporting cells of indifferent gonads of both sexes. In medaka, a mutation in the receptor for Amh, AmhrII, resulted in sex reversal of the gonads in half of the XY fish that had the mutant gene (Morinaga et al., 2007).

3.2. Ovarian Differentiation Ovaries in adult fishes possess oocytes at various stages of development, granulosa cells, theca cells, and blood vessels. During embryogenesis in most fish species, the primordial gonads of genetic males and females are morphologically indistinguishable. In zebrafish, all the hatched fry, regardless of genetic sex, possess gonads with primary oocytes. Later in presumptive testes, the primary oocytes undergo apoptosis, giving way to the development of testicular tissue. On the other hand, in medaka, germ cells start to proliferate and increase in number first in the presumptive ovaries. Germ cell proliferation is followed by entry into a clonal mode of proliferation where dividing germ cells form clusters. These germ cells then enter into meiotic prophase and start to develop as oocytes. Thus, the ovaries in medaka acquire a distinct morphology at very early stages of development. An earlier study postulated that, in mammals, initiation of ovarian formation occurred by default because germ cells that migrated to areas other than gonadal anlagen developed as oocytes (McLaren & Southee, 1997). However, recent evidence has proved that ovarian formation in mammals also is regulated actively at the genetic level. Genes such as Wnt4 (wingless-type MMTV integration site family, member 4) (Kim et al.,

A Y

C

B

X

Cyp19a1a E2

Y

X

Cyp19a1a E2 Exogenous

X

E

D

X

X

X

Cyp19a1a

Cyp19a1a

E2

E2

ER

ER

ER

Testis

Ovary

X

F

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Cyp19a1a E2 Exogenous

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Cyp19a1a E2

E2

E2 ER

2006), Foxl2 (Forkhead box L2) (Ottolenghi et al., 2007; Garcia-Ortiz et al., 2009), and Rspo1 (Roof plate-specific Spondin 1,R-Spondin 1) (Parma et al., 2006) were found to be critical for ovarian differentiation and morphogenesis in mammals. In fishes, there have not been many studies revealing the molecular mechanisms underlying ovarian formation. In several fish species (e.g., tilapia), ovarian differentiation is primarily under the control of E2 (Yamamoto, 1969; Piferrer et al., 1991; Nakamura, Kobayashi, Chang, & Nagahama, 1998; Kobayashi, Kajiura-Kobayashi, & Nagahama, 2003; Guiguen, Fostier, Piferrer, & Chang, 2010) (Figure 1.4). The gene Cyp19 codes for the steroidogenic enzyme, P450aro, responsible for estrogen production in steroidogenic tissues such as the brain, gonads, adrenal gland, etc. In fishes, there are two forms of P450aro, the ovarian form and the brain form, encoded by two different genes, cyp19a1a and cyp19a1b; cyp19a1a is expressed in the ovaries, while cyp19a1b is found predominantly in the brain. Cyp19a1a-expressing cells were apparent near blood vessels in the primordial gonads of tilapia at 7 dah (Sakai, Kobayashi, Matsuda, & Nagahama, 2008). Blocking production of E2 through inhibition of P450aro caused testicular differentiation in XX tilapia fry (Kobayashi, Kajiura-Kobayashi, & Nagahama, 2003), demonstrating that E2 is the natural inducer of ovarian differentiation in this species. More recent investigations in tilapia revealed that cyp19a1a was one of the first genes to exhibit a sexually dimorphic expression pattern prior to any morphological sex differentiation in the primordial gonads (Ijiri et al., 2008) (Figure 1.3). cyp19a1a transcripts were abundant in the XX females of this species at 5 dah, while XY fry of the same stage did not show any such rise in the expression levels of cyp19a1a. Genes coding for other steroidogenic enzymes involved in the E2 pathway are expressed during the same time frame. As expression of cyp19a1a signifies the production of E2, all these data collectively confirm the

ER ER

Ovary

Testis

Ovary

Testis

FIGURE 1.4 17b-estradiol (E2)/estrogen receptor (ER) signaling in ovarian differentiation among fishes. A: ER is present in XY gonads but the absence of Cyp19a1a and E2 primes the gonads to follow the testicular differentiation pathway during normal sexual differentiation. B: when E2 is administered, XY gonads develop as ovaries instead of testes. C: in normally differentiating genetic females, increases in E2 signals, produced by abundant levels of Cyp19a1a, acting through ERs, trigger ovarian development. D: treatment of XX fish with aromatase inhibitors (fadrozole) results in testicular development in the absence of E2/ER signaling. E: administration of E2 rescues the XX gonads in D from undergoing male development. F: finally, blocking ERs with antagonists such as tamoxifen directs the XX gonads toward the male pathway and testicular development.

8

revealed that the role of Foxl2 in ovarian differentiation is conserved among vertebrates. Initiation of foxl2 expression has been linked to ovarian differentiation in a number of teleosts, e.g. tilapia (Wang, Zhou, Kobayashi, & Nagahama, 2004; Wang et al., 2007; Ijiri et al., 2008) (Figure 1.4), medaka (Nakamoto, Matsuda, Wang, Nagahama, & Shibata, 2006; Nakamoto et al., 2009), rainbow trout (Baron et al., 2004; Baron, Houlgatte, Fostier, & Guiguen, 2005), and Silurus meridionalis (Liu, Zhang, & Wang, 2008). foxl2 mRNA was detected specifically in the somatic cells in presumptive ovaries, and overexpression of a dominant negative mutant form of Foxl2 in tilapia demonstrated that this factor was critical for ovarian differentiation and morphogenesis (Wang et al., 2007). Overexpression of foxl2 genomic DNA in genetically male embryos of tilapia resulted in partial to complete sex reversal of the gonads into ovaries. Promoter analysis revealed that Foxl2 regulates the transcription of cyp19a1a either by directly binding to the promoter of the latter or through an interaction with Ad4bp/sf-1 (Figure 1.5). Thus, in tilapia, Foxl2 is essential for ovarian differentiation and morphogenesis. Very recently, conditional ablation of Foxl2 in adult female mice resulted in transdifferentiation of the granulosa cells into Sertoli cells, showing the essential role of FOXL2 in maintaining the ovarian phenotype even after completion of sexual development (Uhlenhaut et al., 2009). This study demonstrated active repression of the male pathway in granulosa cells by FOXL2. Exploration of

20

Relative Luciferase Activity

notion that E2 is critical for female sexual differentiation in tilapia. Further, in rainbow trout, administration of 17aethinylestradiol during the first two months of development upregulated early female genes including cyp19a1a and foxl2, confirming that E2 has a leading role in the induction of ovarian differentiation in this species also (VizzianoCantonnet et al., 2008; Guigen et al., 2010). In zebrafish, cyp19a1a expresses in excess in those gonads that will be transformed into ovaries (Jørgensen et al., 2008). It has been suggested that, in presumptive testes, decrease in cyp19a1a expression leads to apoptosis of the oocytes, eventually resulting in the emergence of testicular tissue (Wang & Orban, 2007). For sex steroids to have their effects during gonadal development, their receptors must also be present. In tilapia, all three types of estrogen receptor (ER) (ERa, ERb1, and ERb2) were consistently expressed at relatively high levels at 5 dah in gonads of both sexes, with no sexually dimorphic expression until 35 dah (Ijiri et al., 2008). Similar levels of expression of ERa in undifferentiated XX and XY gonads were reported in medaka (Kawahara, Omura, Sakai, & Yamashita, 2003) and rainbow trout (Guiguen et al., 1999). These results indicate that E2 synthesized in female gonads mediates female sexual differentiation by stimulating development of undifferentiated XX gonads through ERs. Expression of ERs in XY gonads early during differentiation explains the susceptibility of males to feminization by exogenous E2. Nevertheless, the role of E2 in female sexual differentiation of medaka remains unresolved. Although exogenous E2 administration successfully induces ovarian differentiation in the gonads of genetic males, treatment with the P450aro inhibitor fadrozole during embryogenesis does not cause sex change in genetic females (Kawahara & Yamashita, 2000). Expression of cyp19a1a in medaka is initiated only after morphological sex differentiation has been initiated in the presumptive ovaries. However, in this experiment, the embryos were exposed to fadrozole from the day of fertilization to the day of hatching only and, in medaka, cyp19a1a transcripts are found only from 5 dah (Suzuki, Tanama, Nagahama, & Shibata, 2004). Whether fadrozole remains in the hatched fry even after the withdrawal of the drug was not confirmed in this study. Therefore, it is necessary to ascertain whether extending the treatment until after the initiation of cyp19a1a expression might cause sex reversal. Alternatively, next-generation P450aro inhibitors such as letrozole, anastrozole, and exemestane also could be tried to reverse the phenotypic sex of genetic females. Foxl2 is a transcription factor belonging to the forkhead family. In goats, Foxl2 has been implicated in ovarian differentiation, possibly by regulating the transcription of the CYP19 gene (Pailhoux et al., 2005; Pannetier et al., 2006). Similarly, investigations using teleosts have


Foxl2

Ad4bp/sf-1 cyp19a1a

10

0 Ad4bp/sf-1 Foxl2

100

100 10

50

100

250

100

FIGURE 1.5 The effect of Foxl2 and Ad4bp/sf-1 on the promoter activity of tilapia cyp19a1a in TM3 cells. Foxl2 and Ad4bp/sf-1 expression vectors were cotransfected with tilapia cyp19a1a -2346-bp promoter construct into TM3 cells. Firefly and Renilla luciferase activities were estimated after 48 hours of transfection. Ad4bp/sf-1 or Foxl2 alone increased transcription of cyp19a1a. Foxl2 activated cyp19a1a transcription in a dose-dependent manner. A dramatic rise in the activity of cyp19a1a promoter was apparent when Foxl2 was cotransfected together with Ad4bp/sf-1.

Chapter | 1


a similar role for FOXL2 in ovarian sex differentiation and maintenance of ovarian phenotype among fishes might provide insight into the labile nature of sex determination among fishes. Ad4bp/sf-1 is involved in sexual differentiation and early gonadogenesis in vertebrates. Recently, in mice, association of Sry with SF-1 was found to be critical for testicular differentiation (Sekido & Lovell-Badge, 2008). On the other hand, studies involving the role of Ad4bp/sf-1 role in fish sexual differentiation linked it only with initiation of ovarian differentiation. During early stages of ovarian differentiation in tilapia XX gonads, interaction between Ad4bp/sf-1 and Foxl2 increases the transcription of cyp19a1a, accelerating E2 production in a femalespecific manner (Wang et al., 2007) (Figure 1.4). Foxl2 interacts with the ligand-binding domain of Ad4bp/sf-1 through its forkhead domain to form a heterodimer, which further enhances cyp19a1a transcription, demonstrating that, though ad4bp/sf-1 does not exhibit a sexually dimorphic expression pattern in the gonads during early stages of sexual differentiation, it plays pivotal roles in sex-specific development of the gonads through its association with other sex-specific factors. Further research on the mechanisms involved in testicular differentiation of tilapia revealed that Dmrt1 repressed the female pathway in XY gonads by suppressing the basal as well as the Ad4bp/sf-1activated transcription of cyp19a1a, proving that Ad4bp/ sf-1 is critical for ovarian differentiation of tilapia (Wang et al., 2010). A few studies associate Wnt4 with initiation of ovarian development in fishes. In the protandrous black porgy (Acanthopagrus schlegeli), wnt4 expression is closely linked with ovarian development (Wu & Chang, 2009; Wu et al., 2009). In this species all the fry first develop as males and, after three years, there is male-to-female conversion, during which WNT4, along with Cyp19a1a and Foxl2, is instrumental in the initiation of ovarian tissue development. Nevertheless, the role of Wnt4 in female sex determination and ovarian development among fishes remains largely uncertain.

4. ENVIRONMENTAL EFFECTS ON SEX DETERMINATION AND DIFFERENTIATION Many fish species exhibit ESD. In these fishes, temperature, pH, or social factors determine the developmental fate of the bipotential gonad during the critical period of larval development (Barroiller & D’Cotta, 2001; see also Chapter 8, this volume). Temperature-dependent sex determination (TSD) was reported for the first time in fishes in the Atlantic silverside (Menidia menidia) (Conover & Kynard, 1981). Menidia peninsulae and M. menidia are the only two species that exhibit TSD in the wild (Ospina-Alvarez &

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Piferrer, 2008). The remaining species were tested for TSD only under laboratory conditions and it is still unclear whether they adopt TSD in their natural habitats. In general, three different patterns of sex ratio responses due to thermal influences have been observed among fishes: (1) higher incidence of males at high temperatures, (2) higher incidence of females at low temperatures, and (3) both low and high temperatures resulting in more males, whereas an intermediate temperature produces both males and females in equal proportions (Devlin & Nagahama, 2002; Munch & Conover, 2004). Under laboratory conditions, TSD occurs in cichlids, sea bass (Dicentrarchus labrax), atherinids, and many other teleosts (Ospina-Alvarez & Piferrer, 2008). Thermal influences on sexual differentiation have been demonstrated among teleosts, even in species with GSD (Koshimizu, Stru¨ssman, Okamoto, Fukuda, & Sakamoto, 2010). Gonads of tilapia (as reviewed in Nagahama, Nakamura, Kitano, & Tokumoto, 2004; Baroiller, D’Cotta, & Saillant, 2009) and medaka (Hattori et al., 2007), two species with proven GSD mechanisms, were susceptible to thermal effects during critical periods of ontogenesis. Demonstration of TSD in species with GSD mechanisms poses the question of whether fishes do possess actual TSD mechanisms like reptiles. A reinvestigation to assess the prevalence of TSD among fishes has revealed that TSD is not as widely spread as was previously thought (Ospina-Alvarez & Piferrer, 2008). In this study, 59 species with known TSD mechanisms were rechecked at a range of temperatures during development under natural conditions (range of temperature during development in the wild (RTD)); RTD is more ecologically relevant and, therefore, thermal effects on sex determination should ideally occur in a true TSD species within the range of their RTDs. Using this criterion, it was found that approximately 75% (19 out of 26) of the species reported as having TSD did not actually possess TSD. Instead, sex determination and further sexual differentiation in the gonads of these fishes were the result of thermal effects, especially extreme temperature fluctuations outside their RTDs, on their GSD. These authors concluded that there exists only one general pattern of sex ratio response to temperature, not three as described above, and suggested that fishes in general do not possess a true TSD mechanism, but sexual development in fishes is only sensitive to temperature. Unlike reptiles, where orthologs for most of the factors involved in the mammalian sex determination and differentiation cascade have been found (as reviewed in Georges, Ezaz, Quinn, & Sarre, 2010), information on genetic factors involved in purported TSD of fishes largely remain unknown. The main factor that has been reported to be involved in TSD of fishes is the gene that encodes P450aro (cyp19a1a). At male-producing temperatures, transcripts of cyp19a1a diminish, driving the gonads toward testicular differentiation. Female-producing temperatures cause an

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increase in the expression levels of cyp19a1a, resulting in ovarian differentiation. In the Japanese flounder, high water temperature caused sex reversal from genetic females to phenotypic males and suppression of cyp19a1a expression. Further, follicle-stimulating hormone (FSH) signaling and Foxl2 were implicated in the regulation of cyp19a1a transcription during temperature-dependent sexual determination in this species (T. Yamaguchi, S. Yamaguchi, Hirai, & Kitano, 2007). In the atherinid Odontesthes bonariensis (pejerrey fish), with known TSD, cyp19a1a expression was closely correlated with ovarian formation in fish reared at feminizing temperatures (Fernandino et al., 2008). Conversely, masculinizing temperatures reduced the expression levels of this gene, while mixed-sex-producing temperatures showed a bimodal pattern of expression, indicating the involvement of cyp19a1a in the TSD of this species (Karube et al., 2007). In addition, a rise in dmrt1 expression was correlated with male-producing temperatures (Fernandino et al., 2008; Lee et al., 2009). Exposure to high temperature during early gonadal development in pufferfish (T. rubripes) caused degeneration of germ cells and subsequent masculinization of the somatic cells (Lee et al., 2009). Whether this is a common mechanism during high-temperature-induced testicular differentiation is yet to be ascertained. Regulation of the genes involved in TSD is not well understood. However, a correlation between circulating levels of insulin-like growth factor-1 (IGF-1) and changes in specific growth rate of body size at varying temperatures and different nutritional status has been demonstrated in the southern flounder (Paralichthys lethostigma). Plasma IGF1, muscle IGF-1, and specific growth rate were suppressed at high rearing temperatures relative to low rearing temperatures (Luckenbach et al., 2007). Further analysis of this gene might reveal a possible correlation between TSD and the endocrine-growth regulatory axis. Though pH has been shown to influence sex determination in fishes belonging to the genera Apistogramma and Pelvicachromis (Rubin, 1985; Roëmer & Beisenherz, 1996), sex ratio response to pH is in general less obvious in comparison to temperature. In the molly Peocilia sphenops, a combined effect of pH and temperature was found to skew the sex ratio towards one sex (Baron, Buckle, & Espina, 2002). However, molecular mechanisms underlying pHinduced sex determination have not been studied in detail. Socially controlled sex determination was explained for the first time in the fairy basslet (Gramma loreto) (as reviewed in Devlin & Nagahama, 2002). Social factors were found to play a major role in determining the sex in several teleost hermaphroditic species such as wrasses, damsel fishes, parrotfishes, and gobies (see also Chapter 8, this volume). Relative size and sex ratio are two prominent social factors used among fishes for sex determination and differentiation. In the midas cichlid (Cichlasoma


citrinellum), sex is determined by relative size of an individual during its juvenile stage (Francis & Barlow, 1993). In the protogynous epinepheline Cephalopholis boenak, all isolated, single juveniles differentiated as males, and they clearly showed a faster growth rate than differentiating females and undifferentiated juveniles. In the bluehead wrasse (Thalassoma bifasciatum), population size during juvenile stages has an influence on the development of the primary males (Munday, Wilson, & Warner, 2006). Likewise, in false clown anemonefish (Amphiprion ocellaris) longterm social interactions promote male sexual development and growth, and 11-KT has been implicated as having a role in this process (Iwata, Nagai, Hyoudou, & Sasaki, 2008). The serial sex-changing fish Trimma okinawae possesses both testis and ovary at the same time; however, only one gonad remains fully developed (active) at a time, while the other remains in an underdeveloped state (inactive). According to their social status, these fish are capable of changing their gonadal phenotype back and forth several times. Recent evidence revealed that, soon after the change in the social status of these fish, GTH receptor genes switch their location of expression from the active gonad to the inactive gonad and initiate the development of the inactive gonad (Kobayashi et al., 2009). This fish is a good model for the study of the role of the brain in mediating gonadal sex differentiation in response to environmental stimuli.

5. CONCLUSIONS AND FUTURE DIRECTIONS Genetic and molecular approaches have been used to facilitate the studies in the field of fish sex determination/differentiation. Figure 1.6 summarizes the factors involved in sex determination/differentiation among fishes. Discovery of Dmy from only two species of medaka as the second male sex-determining gene with no structural similarity to the mammalian sex-determining gene, SRY/Sry, has revealed the diversity at the top of the sex-determining cascade among vertebrates. It would be very exciting to see why such a high level of diversity is adopted in the case of sex-determining genes. Fishes might answer this question because they use a wide array of sex-determining mechanisms, ranging from genetic to environmental. More impetus has to be given to studies meant to unveil the targets of Dmy. As with the close association between temporal and spatial expression patterns of dmy and gsdf in medaka, future studies should address the nature of the relationship between these two factors in vivo. The new factor Gsdf is unique to teleosts, suggesting that it plays some role specific to testicular differentiation of this group. Detailed analyses of Gsdf might provide new insights into differences between mammals and fishes, especially with respect to the anatomy of testicular development in these two groups. Another factor crucial for gonadal sex

Chapter | 1

11


GSD

XY Initial trigger

ESD

XX Temperature pH Social factors

Dmy/Dmrt1bY Male pathway

Female pathway

Gsdf

Foxl2

Dmrt1

Ad4bp/sf-1

Sox9a2

Cyp19a1a

Male pathway

Female pathway

Dmrt1

Cyp19a1a

11-KT

E2

FIGURE 1.6 Factors involved in genetic sex determination (GSD) and environmental sex determination (ESD) in teleosts (refer to the text for detailed description). 11-KT, 11-ketotestosterone; E2, 17b-estradiol. See color plate section.

Major player Amh

Hormone

11-KT

E2

differentiation among nonmammalian vertebrates is gonadal sex steroids. It is now widely acknowledged that the estrogen/ER system plays crucial roles in ovarian differentiation in nonmammalian vertebrates, yet the downstream molecular mechanism of estrogen action on ovarian differentiation remains to be determined. Fishes constitute a very good model for studies involving the upstream and downstream molecular cascades leading to the production of E2 and its inductive effects on ovarian development, respectively. In contrast, the possibilities for the involvement of androgens in fish sex determination/differentiation appear to be questionable considering the studies done. The data presented in this review will be helpful in terms of understanding not only the molecular basis of sex determination and differentiation but also the mechanisms underlying sexual plasticity, which fishes exhibit on a large scale. Discovery of more factors and molecular mechanisms involved in sex determination/differentiation would throw some light on the risk factors associated with endocrine disruptors and thus help in addressing the issues associated with the management of potential factors that could be hazardous to the environment. Thus, the next decade might witness radical changes in many central dogmas prevailing in the field of sexual development and plasticity among fishes. This chapter is mainly based on the studies conducted using popular model fishes such as medaka, tilapia, rainbow trout, zebrafish, etc. Data accumulated from these species would be useful in the study of sex determination/differentiation in other teleosts and nonteleosts such as agnathans, condrichthyans, sarcopterygians, chondrosteans, and holosteans. Such studies might reveal the actual degree of diversity in sex determination/differentiation of fishes, especially with respect to the evolutionary mechanisms that

constitute the foundation of vertebrate sex determination and differentiation.

ACKNOWLEDGEMENTS This work was supported in part by grant-in-aid for research from SORST, JST (Japan Science and Technology Corporation) and the Ministry of Education, Science, Sport, and Culture of Japan. The authors declare that there is no conflict of interest that would prejudice the impartiality of this scientific work.

ABBREVIATIONS 11-KT 17b-HSD1 3b-HSD AMH AMHRII BAC CMV CYP11A1 CYP11B2 CYP17a1 CYP19A1 dah dpf E2 EGFP ER ESD Foxl2 FSH gripNA GSD Gsdf

11-ketotestosterone 17b-hydroxysteroid dehydrogenase type 1 3b-hydroxysteroid dehydrogenase Anti-Mu¨llerian hormone Anti-Mu¨llerian hormone receptor 2 Bacterial artificial chromosome Cytomegalovirus See P450scc See P45011b See P45017a See P450aro Days after hatching Days post fertilization 17b-estradiol Enhanced green fluorescent protein Estrogen receptor Environmental sex determination Forkhead box L2 Follicle-stimulating hormone Engineered peptide nucleic acid Genetic sex determination Gonadalesoma-derived factor

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hCREB IGF-1 LG ORF P45011b P45017a P450aro P450scc PCR PGC Rspo1 RTD SD SF-1 T TSD Wnt4


Human cAMP response element-binding Insulin-like growth factor-1 Linkage group Open reading frame 11b-hydroxylase 17a-hydroxylase Ovarian aromatase gene Cholesterol side-chain cleaving enzyme Polymerase chain reaction Primordial germ cell Roof plate-specific Spondin 1,R-Spondin 1 Range of temperature during development in the wild Sex determination Steroidogenic factor-1 Testosterone Temperature-dependent sex determination Wingless-type MMTV integration site family, member 4

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Morinaga, C., Saito, D., Nakamura, S., Sasaki, T., Asakawa, S., Shimizu, N., et al. (2007). The hotei mutation of medaka in the anti-Mu¨llerian hormone receptor causes the dysregulation of germ cell and sexual development. Proc. Natl. Acad. Sci. USA, 104, 9691e9696. Munch, S. B., & Conover, D. O. (2004). Nonlinear growth cost in Menidia menidia: theory and empirical evidence. Evolution., 58, 661e664. Munday, P. L., Wilson, W. J., & Warner, R. R. (2006). A social basis for the development of primary males in a sex-changing fish. Proc. Biol. Sci., 273, 2845e2851. Nagahama, Y., Nakamura, M., Kitano, T., & Tokumoto, T. (2004). Sexual plasticity in fish: a possible target of endocrine disruptor action. Environ. Sci., 11, 73e82. Nakamoto, M., Matsuda, M., Wang, D. S., Nagahama, Y., & Shibata, N. (2006). Molecular cloning and analysis of gonadal expression of Foxl2 in the medaka, Oryzias latipes. Biochem. Biophys. Res. Commun., 344, 353e361. Nakamoto, M., Muramatsu, S., Yoshida, S., Matsuda, M., Nagahama, Y., & Shibata, N. (2009). Gonadal sex differentiation and expression of Sox9a2, Dmrt1, and Foxl2 in Oryzias luzonensis. Genesis, 47, 289e299. Nakamoto, M., Suzuki, A., Matsuda, M., Nagahama, Y., & Shibata, N. (2005). Testicular type Sox9 is not involved in sex determination, but might be in the development of testicular structures in the medaka, Oryzias latipes. Biochem. Biophys. Res. Commun., 333, 729e736. Nakamura, M., Kobayashi, T., Chang, X. T., & Nagahama, Y. (1998). Gonadal sex differentiation in teleost fish. J. Exp. Zool., 281, 362e372. Nakamura, S., Aoki, Y., Saito, D., Kuroki, Y., Fujiyama, A., Naruse, K., et al. (2008). Sox9b/sox9a2-EGFP transgenic medaka reveals the morphological reorganization of the gonads and a common precursor of both the female and male supporting cells. Mol. Reprod. Dev., 75, 472e476. Nanda, I., Kondo, M., Hornung, U., Asakawa, S., Winkler, C., Shimizu, A., et al. (2002). A duplicated copy of FMRT1 in the sexdetermining region of the Y chromosome of the medaka, Oryzias latipes. Proc. Natl. Acad. Sci. U.S.A., 99, 11778e11783. Ospina-Alvarez, N., & Piferrer, F. (2008). Temperature-dependent sex determination in fish revisited: prevalence, a single sex ratio response pattern, and possible effects of climate change. PLoS. One 3, e2837. Otake, H., Masuyama, H., Mashima, Y., Shinomiya, A., Myosho, T., Nagahama, Y. et al. (2009). Heritable artificial sex chromosomes in the medaka, Oryzias latipes. Heredity. In Press. Ottolenghi, C., Pelosi, E., Tran, J., Colombino, M., Douglass, E., Nedorezov, T., et al. (2007). Loss of Wnt4 and Foxl2 leads to femaleto-male sex reversal extending to germ cells. Hum. Mol. Genet., 16, 2795e2804. Pailhoux, E., Vigier, B., Schibler, L., Cribiu, E. P., Cotinot, C., & Vaiman, D. (2005). Positional cloning of the PIS mutation in goats and its impact on understanding mammalian sex-differentiation. Genet. Sel. Evol. 37 Suppl., 1, S55e64. Pannetier, M., Fabre, S., Batista, F., Kocer, A., Renault, L., Jolivet, G., et al. (2006). Foxl2 activates P450 aromatase gene transcription: towards a better characterization of the early steps of mammalian ovarian development. J. Mol. Endocrinol., 36, 399e413. Parma, P., Rado, O., Vidal, V., Chaboissier, M. C., Dellambra, E., Valentini, S., et al. (2006). R-spondin 1 is essential in sex determination, skin differentiation and malignancy. Nat. Genet., 38, 1304e1309.

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Uhlenhaut, N. H., Jakob, S., Anlag, K., Eisenberger, T., Sekido, R., Kress, J., et al. (2009). Somatic sex reprogramming of adult ovaries to testes by Foxl2 ablation. Cell, 139, 1130e1142. Vizziano, D., Randuineau, G., Baron, D., Cauty, C., & Guiguen, Y. (2007). Characterization of early molecular sex differentiation in rainbow trout, Oncorhynchus mykiss. Dev. Dyn., 236, 2198e2206. Vizziano-Cantonnet, D., Baron, D., Mahe´, S., Cauty, C., Fostier, A., & Guiguen, Y. (2008). Estrogen treatment up-regulates but does not suppress all testicular markers during rainbow trout male-to-female gonadal transdifferentiation. J. Mol. Endocrinol., 41, 277e288. Wang, D. S., Kobayashi, T., Zhou, L. Y., Paul-Prasanth, B., Ijiri, S., Sakai, F., et al. (2007). Foxl2 up-regulates aromatase gene transcription female-specifically by binding to the promoter as well as interacting with Ad4BP/SF-1. Mol. Endocrinol., 21, 712e725. Wang, D. S., Zhou, L. Y., Kobayashi, T., Matsuda, M., Shibata, S., Sakai, F., et al. (2010). Dmrt1 repression of aromatase transcription, a possible mechanism favoring the male pathway in tilapia. Endocrinology, 151, 1331e1340. Wang, D. S., Zhou, L. Y., Kobayashi, T., & Nagahama, Y. (2004). Molecular cloning and gene expression of Foxl2 in the Nile tilapia, Oreochromis niloticus. Biochem. Biophys. Res. Commun., 320, 83e89. Wang, X. G., & Orban, L. (2007). Anti-Mu¨llerian hormone and 11 betahydroxylase show reciprocal expression to that of aromatase in the transforming gonad of zebrafish males. Dev. Dyn., 236, 1329e1338. Wilhelm, D., Washburn, L. L., Truong, V., Fellous, M., Eicher, E. M., & Koopman, P. (2009). Antagonism of the testis- and ovary-determining pathways during ovotestis development in mice. Mech. Dev., 126, 324e336. Woram, R. A., Gharbi, K., Sakamoto, T., Hoyheim, B., Holm, L. E., Naish, K., et al. (2003). Comparative genome analysis of the primary sex-determining locus in salmonid fishes. Genome Res., 13, 272e280. Wu, G. C., & Chang, C. F. (2009). Wnt4 associated with the development of ovarian tissue in the protandrous black Porgy, Acanthopagrus schlegeli. Biol. Reprod., 81, 1073e1082. Wu, G. C., Tomy, S., Lee, M. F., Lee, Y. H., Yueh, W. S., Lin, C. J., et al. (2009). Sex differentiation and sex change in the protandrous black porgy, Acanthopagrus schlegeli. Gen. Comp. Endocrinol. doi: 10.1016/j.ygcen.2009.11.003. Yamaguchi, T., Yamaguchi, S., Hirai, T., & Kitano, T. (2007). Folliclestimulating hormone signaling and Foxl2 are involved in transcriptional regulation of aromatase gene during gonadal sex differentiation in Japanse flounder, Paralichthys olivaceus. Biochem. Biophys. Res. Commun., 359, 935e940. Yamamoto, T. (1969). Sex differentiation. In W. Hoar, & D. Randall (Eds.), Fish Physiology, Vol. III (pp. 117e175). Academic Press. Yoshimoto, S., Okada, E., Umemoto, H., Tamura, K., Uno, Y., NishidaUmehara, C., et al. (2008). A W-linked DM-domain gene, DM-W, participates in primary ovary development in Xenopus laevis. Proc. Natl. Acad. Sci. USA, 105, 2469e2474. Yoshinaga, N., Shiraishi, E., Yamamoto, T., Iguchi, T., Abe, S., Kitano, T., et al. (2004). Sexually dimorphic expression of a teleost homologue of Mu¨llerian inhibiting substance during gonadal sex differentiation in Japanese flounder, Paralichthys olivaceus. Biochem. Biophys. Res. Commun., 322, 508e513. Zhou, R., Liu, L., Guo, Y., Yu, H., Cheng, H., Huang, X., et al. (2003). Similar gene structure of two Sox9a genes and their expression patterns during gonadal differentiation in a teleost fish, rice field eel (Monopterus albus). Mol. Reprod. Dev., 66, 211e217.

Chapter 2

Conserved and Divergent Features of Reproductive Neuroendocrinology in Teleost Fishes Olivier Kah* and Sylvie Dufoury *

Universite´ de Rennes, Rennes, France, y Museúm National d’Histoire Naturelle, Paris, France

SUMMARY This chapter will deal mainly with teleost fishes, which constitute the largest (close to 30 000 species) and best-studied group. Despite their phylogenetic distance, fishes share with tetrapods a number of common characteristics with respect to neuroendocrine control of the reproductive axis. Fishes synthesize luteinizing hormone (LH) and follicle-stimulating hormone (FSH), which regulate early gametogenesis, steroidogenesis, and ovulation/ spermiation. Pituitary gonadotropic secretion is regulated by gonadotropin-releasing hormone (GnRH), which is in turn controlled by a number of neurotransmitters and neuropeptides, including kisspeptins (Kps). Gonadal steroids also act at the hypothalamus to regulate secretion of gonadotropins (GTHs). However, due to their long evolutionary history and phylogenetic diversity, many fish species have developed adaptive mechanisms that may differ from one species to the next. Of particular importance in fishes is the role of the environment in sexual differentiation and the diversity of reproductive strategies, leading to multiple variations in a common general mechanism.

1. INTRODUCTION Actinopterygian fishes, also called ray-finned fishes, diverged from sarcopterygians, the lobe-fined fishes, some 450 millions years before present (MYBP). Although sarcopterygian fishes gave rise to land vertebrates, all actinopterygian fishes remained dependent on the aquatic environment and gave birth to the large group of teleost fishes. This group, believed to have emerged 300e350 MYBP, has been extremely successful and currently is comprised of approximately 28 000 species. It is believed that one of the reasons for their evolutionary success is the fact that a third entire genome duplication (often named 3R) occurred early after the teleost emergence, making duplicate sequences available for the Hormones and Reproduction of Vertebrates, Volume 1dFishes Copyright Ó 2011 Elsevier Inc. All rights reserved.

evolution of new functions (Steinke, Hoegg, Brinkmann, & Meyer, 2006). In addition, teleost genomes underwent frequent gene-linkage disruptions compared to other vertebrates. Thus, in teleosts, protein-coding sequences evolved faster than in other groups, also favoring rapid radiation and speciation (Ravi & Venkatesh, 2008). Another reason for this evolutionary success resides in the fact that teleost fishes possess a remarkable capacity to develop unique reproductive strategies, most likely due to an astonishing sexual plasticity at both brain and gonadal levels. Despite the phylogenetic distance between fishes and mammals, and the unique variety of reproductive strategies of fishes (Jalabert, 2005), the basic components of the reproductive axis are relatively similar in all vertebrates. The vertebrate neuroendocrine system (Figure 2.1) consists of neurosecretory neurons located in special nuclei within the hypothalamus of the brain and the hypophysis or pituitary gland (see Norris, 2007). The vertebrate pituitary is separable into the glandular adenohypophysis and the neurohemal neurohypophysis. The adenohypophysis consists of a pars distalis (PD), a pars intermedia (PI), and a pars tuberalis (PT). The neurohypophysis subdivision of the pituitary is separable into two distinct neurohemal areas: (1) the median eminence (ME), connected to the adenohypophysis by the hypothalamusepituitary portal system of blood vessels, and (2) the pars nervosa (PN). Teleost fishes are unique among vertebrates in that the ME is lacking and axons from some of the neurosecretory neurons in the hypothalamus directly innervate the PD and PI (see Zohar, Munoz-Cueto, Elizur, & Kah, 2010). The PT is absent in all fishes and is found only in tetrapod vertebrates. Gonadotropins (GTHs) of fishes share many structural and functional characteristics with their mammalian counterparts. Fish FSH is believed to stimulate follicular 15

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FIGURE 2.1 (a and b) Comparative organization of the hypothalamoepituitary connections in teleost fishes (a) and mammals (b). In fishes, there is no hypothalamusepituitary portal system. The hypophysiotropiceneurons send their projections directly into the anterior lobe. Nerve endings either connect with secretory cells of the anterior lobe (1a; see (c) and (d)) or end at a basal membrane that separates the neurohypophysis (nh) from the adenohypophysis (1b). This innervation corresponds to that of the median eminence in mammals (1). In both fishes and mammals, magnocellular neurons (oxytocin/isotocin and vasopressin/ vasotocin) send their projections to the neural lobe (¼ pars nervosa), which in teleost fishes is intermingled with the pars intermedia. (c and d) Examples of g-aminobutyric acid (GABA) terminals directly apposed to LHb (gonadotropin (GTH)) cells (a) or growth hormone (GH) cells (b). gaminobutyric acid fibers were labeled by a preembedding technique using peroxidase, while pituitary hormones were labeled by postembedding immunohistochemistry with gold particles. Bar ¼ 0.5 mm. Reproduced from Kah et al. (1992), with permission from S. Karger AG, Basel. See color plate section.

growth in the ovary and spermatogenesis in the testis, whereas LH is involved in the control of the final steps leading to ovulation and spermiation (Levavi-Sivan, Bogerd, Manãno´s, Go´mez, & Lareyre, 2010). Fishes and tetrapods share many of the mechanisms controlling GTH secretion. This chapter summarizes the most recent thinking on the reproductive neuroendocrinology of teleosts, emphasizing both conserved and divergent features.

2. THE UNIQUE HYPOTHALAMICe PITUITARYeGONADAL (HPG) AXIS OF TELEOSTS The evolutionary origin of the pituitary is unclear as it is found only in craniates. Although some immunocytochemical studies have targeted Hatschek’s pit, found in cephalochordates, as a putative progenitor of the vertebrate pituitary, this matter is far from settled (see Christiaen

et al., 2002; Jaszczyszyn et al., 2007; Joly et al., 2007). Recent data have identified proteins in vertebrate and protostome genomes, structurally related to glycoprotein hormone subunits, named glycoproteins a2 (GPA2) and b5 (GPB5) (Hsu, Nakabayashi, & Bhalla, 2002). Recombinant human GPA2/GPB5 heterodimer was shown to activate the thyrotropin (TSH) receptor and was therefore named ‘thyrostimulin’ (Nakabayashi et al., 2002). However, the physiological roles of GPA2 and GPB5 are still unknown. Spatiotemporal expression of GPA2 and GPB5 at different developmental stages in a basal chordate (amphioxus (Branchiostoma lanceolatum)) suggests that these subunits may play differential roles during embryogenesis without being obligatorily dimerized (Dos Santos et al., 2009). Altogether these data indicate that ancestors of the genes encoding pituitary hormones can be found in invertebrates. In any case, the earliest craniates such as the jawless fishes (Agnatha) already have a well-differentiated pituitary that secretes virtually all of the classical pituitary hormones

Chapter | 2

Conserved and Divergent Features of Reproductive Neuroendocrinology in Teleost Fishes

(Uchida, Murakami, Kuraku, Hirano, & Kuratani, 2003; Kawauchi & Sower, 2006; Sower, Freamat, & Kavanaugh, 2009). This means that we lack some key intermediate species that would fill the huge gap in our understanding of the evolution of the pituitary. This implies that early craniates already had developed in parallel some sort of pituitary gland and the neuroendocrine systems for controlling its main functions. This also means that many of the molecular and cellular mechanisms that were recruited for control of the pituitary probably pre-existed in invertebrates. In that respect, the well-documented example of gonadotropin-releasing hormone (GnRH), which seems to be a heritage from invertebrates, is quite interesting (Twan et al., 2006; Kah et al., 2007; Okubo & Nagahama, 2008; Sower et al., 2009). According to this reasoning, it is not surprising that many vertebrate neuroendocrine factors also exist in invertebrates. It is thus likely that neuronal systems making up these factors were progressively recruited to control new intermediate endocrine glands serving an amplification purpose. Indeed, the ‘invention’ of pituitary hormones allowed a small number of neurons synthesizing minute quantities of neuropeptides to exquisitely control the release, and in some cases the synthesis, of hormones massively liberated into the blood to influence distant glands. A unique feature of the teleost pituitary is the absence of a portal blood connection and its direct innervation by hypothalamic neurons (Figure 2.1), although blood vessels passing from the ventral hypothalamus to the pituitary have been reported in goldfish (Carassius auratus) (Richard E. Peter, personal communication). It is believed that the absence of a blood connection in teleosts is a secondary characteristic because an ultra-short vascular network, resembling the hypothalamusepituitary portal system, has been reported in a more primitive, nonteleost actinopterygian, the Caspian sturgeon (Acipenser guldenstadti) (Polenov & Pavlovic, 1978). For example, no GnRH fibers were observed in the anterior lobe of the sturgeon pituitary. In contrast, they seem to end in the basal hypothalamus just above the anterior lobe of the pituitary (Lepreˆtre et al., 1993). Lampreys also lack direct pituitary innervation and have a poorly described zone of diffusion between the ventral hypothalamus and the pituitary (Polenov, Belenky, & Konstantinova, 1974; Tsuneki & Gorbman, 1975; Sower & Kawauchi, 2001). In teleosts, the neurohormones controlling the activity of the different cell types are released directly by nerve endings located in close vicinity of their target cells (Figure 2.1). In some species, true neuroendocrine synaptic contacts can even be observed (Kaul & Vollrath, 1974; Zohar et al., 2010). The pituitary innervation of teleosts must thus be considered as the functional equivalent of the ME in chondrichthyan and more primitive bony fishes as

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well as in tetrapods (Figure 2.1(b)). In some species, such as salmonids or eels, the neurohypophysis is totally separated from the adenohypophysis by a double basal membrane on which the neurosecretory terminals are apposed (Abraham, 1974; Kaul & Vollrath, 1974). It is believed that the neurosecretory products are released and diffuse within the intracellular spaces to reach their target cells. In other species, such as cyprinids, the basal membrane is interrupted in some places and the nerve fibers invade the adenohypophysis (Figure 2.1(ced)). In that case, it is possible to observe synaptic-like figures with membrane thickening and accumulation of small clear vesicles, strongly suggesting that neurosecretory material is released at these places (Kah et al., 1992; Zohar et al., 2010). In general, pituitary cells of a given type are regionalized in the adenohypophysis of fishes. Lactotropes, which secrete prolactin (PRL), and corticotropes, which secrete corticotropin (ACTH), are usually located in the anterior portion (rostral PD (RPD)); gonadotropes, which secrete FSH and LH, and somatotropes, which secrete growth hormone (GH), occur in the median aspect of the adenohypophysis (proximal PD (PPD)). In addition, there is good correspondence between the distribution of the nerve fibers involved in the control of a given cell type and the distribution of this cell type within the pituitary. For example, GnRH-immunoreactive fibers are mostly present in the PPD, where the gonadotropes are located. Many neuropeptides and neurotransmitters have been identified by immunohistochemistry in fibers penetrating the PD and shown to modulate GTH in release in vivo or in vitro (Kah et al., 1989; Batten, Cambre, Moons, & Vandesande, 1990). This is true for GnRH (Kah et al., 1986; Amano et al., 1991; Gonzalez-Martinez et al., 2002), neuropeptide Y (NPY) (Kah et al., 1989; Batten et al., 1990), dopamine (DA) (Kah, Dubourg, Chambolle, & Calas, 1984; Kah et al., 1986), and g-aminobutyric acid (GABA) (Kah, Dulka, Dubourg, Thibault, & Peter, 1987) (Figure 2.1(ced)). The functions of neurohormones on pituitary activities in teleosts are in many instances not strongly established, and we will restrict ourselves to those clearly serving reproductive functions, namely GnRH, DA, kisspeptins (Kp), GABA, and NPY.

3. GONADOTROPIN-RELEASING HORMONE (GnRH) As in other craniates, GnRH is a key player in the control of the HPG axis in fishes and, because of its use in fish farming, it has received considerable attention. In addition, because commercially relevant species differ from one part of the world to another, numerous species have been studied. In fact, there is no other vertebrate group in which there are so many data available on so many species.

18


3.1. Discovery of Gonadotropin-releasing Hormone (GnRH) in Teleosts There is now strong evidence that GnRH-like peptides existed some 600 MYBP, and appear with their corresponding receptors unambiguously in several nonchordate invertebrate groups as well as in protochordates, where they sustain functions clearly linked to reproductive and some ‘nonreproductive’ functions (Kah et al., 2007). Three GnRH variants have been found in lampreys (lGnRH-I, lGnRH-II, and lGnRH-III; see Table 2.1), whereas only two forms have been reported basal actinopterygians: reedfish (Calamoichthys calabaricus), sturgeon (Acipenser transmontanus), and alligator gar (Lepisosteus spatula) (Sherwood, Doroshov, & Lance, 1991). These two variants correspond to mammalian (mGnRH) and chicken GnRH-II (cGnRH-II) (see Table 2.1), which are believed to be ancestral forms resulting from a previous gene duplication. The first evidence that GnRH exists in teleost fish came from studies showing that sheep brain extracts could stimulate LH release in carp and vice versa (Breton, Jalabert, Billard, & Weil, 1971) and that mGnRH was capable of stimulating gonadotrope activity in carp (Cyprinus carpio) (Breton, Weil, Jalabert, & Billard, 1972). Following these pioneering experiments, numerous

studies examined GnRH effects in many fish species. The first unique teleost GnRH characterized in a teleost fish was purified from a salmon and thus named salmon GnRH (sGnRH) (Sherwood et al., 1983), a decapeptide that differs from mGnRH only by two residues in positions seven and eight (Table 2.1). From early immunohistochemical studies using partially characterized sGnRH antibodies, it appeared that GnRH neurons were distributed along a continuum extending from the olfactory nerve and bulbs to the hypothalamus through the ventral telencephalon and the preoptic area (POA) (Kah et al., 1986). Extending from these cell bodies, immunoreactive (ir) GnRH fibers were identified, coursing towards the pituitary. Teleosts also possess an additional population of large ir-GnRH neurons in the dorsal tegmentum of the midbrain (Kah et al., 1986). Additionally, using specific antibodies, pioneering studies indicated in masu salmon (Onchorynchus masou) that large neurons of the ventral tegmentum actually express cGnRH-II (Amano et al., 1991). It is now established that this second neuronal system synthesizing cGnRH-II in the dorsal tegmentum of the midbrain exists in all teleosts examined and in most vertebrate species (Lethimonier, Madigou, Munoz-Cueto, Lareyre, & Kah, 2004; Guilgur, Moncaut, Canario, & Somoza, 2006; Kah et al., 2007).

TABLE 2.1 Primary structure of the known gonadotropin-releasing hormone (GnRH) variants in craniates. Mammalian GnRH is taken as the reference. The eight variants found in teleosts are marked in bold 1

2

3

4

5

6

7

8

9

10

Reference

Mammalian (mGnRH)

pGlu

His

Trp

Ser

Tyr

Gly

Leu

Arg

Pro

Gly-NH2

Matsuo et al. (1971); Burgus et al. (1972)

Chicken I (cGnRH-I)

-

-

-

-

-

-

Gln

-

-

Miyamoto et al. (1982)

Frog GnRH (frGnRH)

-

-

-

-

-

-

Trp

-

-

Yoo et al. (2000)

Seabream (GnRH)

-

-

-

-

-

-

Ser

-

-

Powell et al. (1994)

Salmon (sGnRH)

-

-

-

-

-

Trp

Leu

-

-

Sherwood et al. (1983)

Whitefish (whGnRH)

-

-

-

-

-

-

Met

Asn

-

-

Adams et al. (2002)

Guinea pig (gpGnRH)

-

Tyr

-

-

-

-

Val

-

-

-

Jimenez-Linãn et al. (1997)

Medaka (mdGnRH)

-

-

-

-

Phe

-

-

Ser

-

-

Okubo et al. (2000)

Chicken II (cGnRH-II)

-

-

-

-

His

-

Trp

Tyr

-

-

Miyamoto et al. (1984)

Catfish (cfGnRH)

-

-

-

-

His

-

-

Asn

-

-

Ngamvongchon et al. (1992)

Herring (hgGnRH)

-

-

-

-

His

-

-

Ser

-

-

Carolsfeld et al. (2000)

Dogfish (dfGnRH)

-

-

-

-

His

-

Trp

Leu

-

-

Lovejoy et al. (1992)

Lamprey I (lGnRH-I)

-

-

-

-

His

Asp

Trp

Lys

-

-

Sherwood et al. (1986)

Lamprey II (lGnRH-II)

-

-

-

-

His

-

Trp

Phe

-

-

Kavanaugh et al. (2008)

Lamprey III (lGnRH-III)

-

-

Tyr

-

Leu

Glu

Trp

Lys

-

-

Sower et al. (1993)

-

Chapter | 2


Thus, in general, the organization of the GnRH systems in teleosts matches that observed in other vertebrates. A first GnRH (type 1) system develops from the olfactory placode and consists of migrated cell bodies dispersed in the ventral forebrain, specifically the subpallium, POA, and anterior mediobasal hypothalamus (MBH). These cells project principally towards the ME in tetrapods or directly into the pituitary, as in teleosts. A second (type 2) GnRH system arises from a mesencephalic primordium. Large neurons synthesizing cGnRH-II project to many brain regions (Lethimonier et al., 2004; Kah et al., 2007).

3.2. Two or Three Gonadotropin-releasing Hormone (GnRH) Genes in Fishes Subsequent studies have identified additional GnRH variants in teleost fishes. Among vertebrates, teleost fishes represent the group with the greatest number of GnRH variants. Indeed, following the identification of sGnRH (Sherwood et al., 1983), seven other GnRH forms have been purified and sequenced in teleosts (Adams et al., 2002), including the mGnRH (King, Dufour, Fontaine, & Millar, 1990) and the evolutionarily conserved cGnRH-II, found in the brains of almost all vertebrates. Many fish species possess up to three GnRH forms generated by three different GnRH genes, as first shown in the African cichlid (Haplochromis burtoni) (White, Kasten, Bond, Adelman, & Fernald, 1995) and gilthead seabream (Sparus auratus) (Gothilf, Elizur, Chow, Chen, & Zohar, 1995) and then in many other species. As indicated in Table 2.1, GnRH variants have been named according to the species in which they were first encountered, but subsequent analyses indicate that GnRH variants in vertebrates are produced from three different paralogous genes: gnrh1, gnrh2, and gnrh3 (Fernald & White, 1999). These GnRH genes encode GnRH preprohormones, the structure of which always consists of a signal peptide along with a prohormone composed of the GnRH decapeptided processing peptide (GKR)dand a GnRH-associated peptide (GAP). A new nomenclature, based on these phylogenetic analyses but also on neuroanatomical and functional considerations, was introduced by Fernald and White (1999). According to this nomenclature, all cGnRH-II (GnRH2) precursors derive from a gnrh2 gene, all sGnRH (GnRH3) precursors derive from a gnrh3 gene, and all other variants (GnRH1) are generated by a gnrh1 gene. Figure 2.2(a) depicts the current hypothesis explaining the relationships among these three paralogous lineages (Guilgur et al., 2006; Kah et al., 2007). This hypothesis postulates that, for invertebrates having at least one GnRH gene, one would expect that the 1R and 2R duplications (Steinke et al., 2006; Kuraku, Meyer, & Kuratani, 2009) gave rise to four

19

potential GnRH genes in basal vertebrates. However, the current information suggests that one copy of these ancestral duplicated genes was rapidly lost. Thus, the current available information suggests that basal representatives of both early actinopterygians and sarcopterygians have a gnrh1 and a gnrh2 gene. Following the teleost-specific genome duplication (Steinke et al., 2006; Kuraku et al., 2009), a second copy of the gnrh1 gene emerged: gnrh3. With respect to the gnrh2 gene, it is probable that the second copy was rapidly lost (Figure 2.2(a)). Another hypothesis, based on syntheny, stipulates that the duplication of gnrh1 occurred before the divergence of the actinopterygians and sarcopterygians, but that this latter branch lost the copy soon after the duplication event (Okubo & Nagahama, 2008). This hypothesis is very possible, but implies that the second copies of all three genes resulting from the teleost-specific gene duplication would have been lost very rapidly after the event.

3.3. Gonadotropin-releasing Hormone (GnRH) Distribution in the Brain of Fishes Three GnRH forms exist in most teleost orders, shown either by cDNA sequencing or biochemical characterization (Carolsfeld et al., 2000; Okubo, Amano, Yoshiura, Suetake, & Aida, 2000; Andersson et al., 2001; Montaner et al., 2001; Adams et al., 2002; Amano, Oka, Yamanome, Okuzawa, & Yamamori, 2002; Mohamed & Khan, 2006). Using specific GAP riboprobes and antibodies, it was found that GnRH2 cells are restricted to the dorsal synencephalon but that GnRH1 and GnRH3 cell distribution overlaps in the olfactory bulbs, ventral telencephalon, and POA (Gonzalez-Martinez et al., 2001; 2002). Similar results were later obtained in other fish species including lake whitefish (Coregonus clupeaformis) (Vickers, Laberge, Adams, Hara, & Sherwood, 2004), a cichlid (Cichlasoma dimerus) (Pandolfi et al., 2005), the Atlantic croaker (Micropogonias undulatus) (Mohamed, Thomas, & Khan, 2005), and medaka (Oryzias latipes) (Okubo et al., 2006), suggesting that this overlap is more the rule than the exception. However, during the early diversification of teleosts, it appears that some groups lost either gnrh1 (Figure 2.2(a)) (salmonids, cyprinids) or gnrh3 (eels, catfishes) (Guilgur et al., 2006; Kah et al., 2007). However, in all cases, the overall distribution of the GnRH neurons remains similar (Figure 2.3). For example, in zebrafish (Danio rerio), a species expressing gnrh2 and gnrh3, gnrh3 neurons are present in regions occupied by both GnRH3 and GnRH1 cells in perciforms (Abraham, Palevitch, Gothilf, & Zohar, 2009) (see Figure 2.3). Studies performed in sea bass (Dicentrarchus labrax) and based on the use of antibodies against GAP, provide unambiguous and detailed information on the distribution

20


FIGURE 2.2 (a) Hypothesis on the evolution of gonadotropin-releasing hormone (GnRH) genes in vertebrates based on the existence of a third full genome duplication, specific to teleost fishes (3R). This hypothesis postulates that invertebrates have at least one gnrh gene, leading to the expectation that the 1R and 2R duplications would have given birth to four potential gnrh genes, as is the case for GnRH receptor (GnRH-R) (see (b)). However, the current information suggests that one copy of these ancestral duplicated genes was rapidly lost. Thus, the current available information suggests that two gnrh genes, gnrh1 (green) and gnrh2 (purple), were present in basal representatives of both early actinopterygians and sarcopterygians. Following the teleost-specific genome duplication, a second copy of the gnrh1 gene emerged (gnrh3, red). However, in some teleost species, either gnrh1 or gnrh3 were lost. With respect to gnrh2, it is probable that the second copy gene was rapidly lost. (b) Hypothesis on the evolution of GnRH-Rs in vertebrates. Starting with one putative ancestral GnRH-R sequence in invertebrates, the 1R and 2R genome duplications potentially generated four sequences in basal sarcopterygians and actinopterygians. In support of this hypothesis, three GnRH-Rs were found in the bullfrog, a diploid species. In teleost fishes, the 3R teleost-specific genome duplication could have generated up to eight putative sequences; accordingly, five functional GnRH-R sequences have been demonstrated in certain teleost species. This indicates that other GnRH-R copies were lost along the way. Reproduced from Kah et al. (2007), with permission from Elsevier. See color plate section.

of ir-GnRH fibers for the three different GnRH forms expressed in the brain of a single species (GonzalezMartinez et al., 2002). Thus, ir-GnRH1 fibers were observed only in the ventral surface of the forebrain (associated with the ventral telencephalon), POA, and hypothalamus, whereas ir-GnRH2 and ir-GnRH3 fibers exhibited a very wide distribution in the brain of the sea bass. Only ir-GnRH1 neurons project to the pituitary, where a dense innervation is observed in the PPD and the periphery of the PI, where gonadotropes with GnRH receptors occur (Gonzalez-Martinez et al., 2004b). This result corroborates physiological evidence suggesting

a major role for GnRH1 in the stimulation of GTH secretion in perciform species. Although ir-GnRH3 axons also reach the pituitary of sea bass, this innervation is much less intense than that of GnRH1. In contrast, no prepro-GnRH2 axons were detected in the pituitary of sea bass, suggesting that the putative role of GnRH2 in the control of reproduction does not involve a direct action of cerebral GnRH2 on gonadotropes, at least in this species. Axonal endings of GnRH-II neurons innervating the pituitary could, however, be detected in other species, such as the European eel (Anguilla anguilla) (Montero, Le Belle, King, Millar, & Dufour, 1995) and goldfish (Kim et al., 1995).

Chapter | 2


21

terminal-nerve neurons that project in many brain regions and where GnRH3 would act as a neuromodulator. A possible new function for GnRH2 has emerged recently from studies in sea bass. In this species, it was shown that GnRH2 neurons in the synencephalon send projections to the pineal gland, thus stimulating melatonin release. This was corroborated by the demonstration of a strong expression of a GnRH-R having strong affinity for GnRH2 (Servili et al., 2010). Figure 2.2(b) proposes a hypothesis for the evolution of GnRH-Rs in vertebrates, notably in fishes. FIGURE 2.3 Organization of the gonadotropin-releasing hormone (GnRH) systems in teleosts. The European eel and African catfish (a) illustrate the situation in which only gnrh1 GnRH is found together with gnrh2 (Dubois, Zandbergen, Peute, Bogerd, & Goos, 2001). In evolved salmonids and cyprinids (b), only gnrh3 is expressed in neurons of the anterior ventral brain. The lake whitefish or modern teleosts (sea bass) (c) are examples of species with three GnRH variants (gnrh1, gnrh2, gnrh3). As one can see, the overall anatomical pattern of expression is similar in all species. Adapted from Lethimonier, Madigou, Munoz-Cueto, Lareyre, and Kah (2004), with permission from Elsevier.

3.4. Gonadotropin-releasing Hormone (GnRH)’s Functions are Mediated by Multiple Receptors The main function of GnRH in fishes is to control the synthesis and release of GTHs (Zohar et al., 2010). However, the presence of three GnRH forms raises questions regarding their respective functions, and the issue is made even more complex because of the multiplicity of GnRH receptors (GnRH-Rs). Five GnRH-R subtypes have been characterized in the sea bass and the fugu (Takifugu rubripes) (Gonzalez-Martinez et al., 2004a; Lethimonier et al., 2004; Moncaut, Somoza, Power, & Canario, 2005), but their presence in other teleost orders is not established (Figure 2.2(b)). The precise distribution and functions of these receptors are still vague and remain open to speculation. In the sea bass, one of these receptors is strongly expressed in the pituitary, while the others are present in multiple organs (Gonzalez-Martinez et al., 2004a; Lethimonier et al., 2004). The GnRH receptor in the pituitary mediates the response to GnRH1 or GnRH3 (when GnRH1 is lacking). Although GnRH3 axons also reach the pituitary of sea bass, this innervation is strongly reduced compared to GnRH1 projections (Gonzalez-Martinez et al., 2002). However, this does not exclude specific functions of GnRH3 in the pituitary, possibly during development or at certain specific physiological stages. In addition, many brain regions exhibit a dense innervation by GnRH2 or GnRH3 fibers. According to detailed studies in sea bass, GnRH3 innervates mainly sensory or neurosensory regions, while GnRH2 is more abundant in sensorimotor regions. The GnRH3 innervation mainly originates from the

3.5. Pulsatile Release of Gonadotropinreleasing Hormone (GnRH) in Fishes? A key feature of the GnRH system in mammals is the pulsatile release of GnRHI from the terminals in the ME. Upon reaching the PD, the pulsatile GnRH signal is translated into a pulsatile release of FSH or LH. The pulsatility of GnRH release is crucial for the regulation of gonadal activity as GnRH not only stimulates the liberation of GTHs, but also their synthesis. Depending on the frequency of the GnRH pulses, the synthesis of FSH (low GnRH frequency) or LH (high GnRH frequency) is favored. This is permitted by activation of a variety of signaling pathways upon activation of a unique G-protein-coupled receptor (GPCR). The mammalian type 1 GnRH-R is the only GPCR lacking an intracellular C-terminal tail. All other GnRH-Rs in vertebrates have an intracellular C tail (Kah et al., 2007). This C tail is essential for homologous desensitization through phosphorylation and b-arrestin-mediated internalization. As a result, the mammalian GnRH receptor does not internalize, but desensitizes according to Ca2þ-dependent mechanisms. For the moment, we have no indication that GnRH release in the pituitary is pulsatile in fishes. The fact that the ME is lacking precludes measurement of GnRH in the portal blood in a similar way to what has been done in mammals (Caraty, Orgeur, & Thiery, 1982). In teleosts, the GnRH peptides accumulate in terminals in the distal neurohypophysis and/or the adenohypophysis and are released upon stimulation. Another potential difference between fishes and mammals concerns estrogen-positive and -negative feedbacks, which are crucial in the regulation of GnRH pulse intervals. These differences make it unlikely that a GnRH pulsatile release in teleosts could be as acurately regulated as it is in mammals. Accordingly, sustained treatments with GnRH (such as implants) are able to activate gonadotropic function in immature teleosts (Crim, 1991; Vidal et al., 2004), whereas this would be achieved only by pulsatile treatments in mammals. In addition, there is evidence that the gonadotropes in teleost fishes receive multiple inputs from a variety of peptidergic, aminergic, and GABAergic

22


FIGURE 2.4 Summary of the differences in the control of gonadotropin release between fishes (a) and mammals (b). The gonadotropes in fishes are known to receive multiple inputs and to carry a large variety of receptors (see Trudeau, 1997; Zohar et al., 2009). These multiple neuronal systems are in turn controlled by the peripheral hormones. In contrast, in mammals the gonadotropin-releasing hormone (GnRH) neuron is the final integrator, the activity of which is regulated by several neurotransmitters and peptide-releasing neurons, which in turn are controlled by estrogens (Moenter, Chu, & Christian, 2009). 5-HT, serotonin; FSH, follicle-stimulating hormone; GABA, g-aminobutyric acid; KISS, kisspeptin; LH, luteinizing hormone; NE, norepinephrine; NPY, neuropeptide Y; VIP, vasoactive intestinal peptide.

neurons (see Section 4). Thus, it seems that the fine tuning of GTH release also takes place at the gonadotrope level in fishes and not mainly at the level of the GnRH neurons as is the case in mammals (Figure 2.4). The possible occurrence of GnRH pulsatility in teleosts deserves further investigation. Electrophysiological studies of the terminal nerve-GnRH neurons in the dwarf gourami (Colisa lalia) revealed a pacemaker activity. Further, the frequency of GnRH pacemaker activity could be modulated by GnRH peptide itself, a mechanism similar to that described for GnRH neurons in mammals (Abe & Oka, 2002).

4. OTHER BRAIN FACTORS STIMULATING GTH RELEASE Apart from GnRH, a large number of neurotransmitters and neuropeptides have been shown to stimulate GTH release directly at the level of the pituitary (Trudeau, 1997; Zohar et al., 2010). Therefore, the regulation of GTH release directly at the level of the pituitary is clearly multifactorial in teleosts (Figure 2.3). Such effects have been reported mainly in goldfish (Somoza & Peter, 1991; Sloley, Kah, Trudeau, Dulka, & Peter, 1992; Himick, Golosinski, Jonsson, & Peter, 1993; Himick & Peter, 1995; Rao, Murthy, Cook, & Peter, 1996; Lin & Peter, 1997; Unniappan et al., 2002; Unniappan & Peter, 2004; Canosa, Stacey, & Peter, 2008).

4.1. Neuropeptide Y (NPY) Neuropeptide tyrosine is a 36-amino-acid peptide of the pancreatic peptide family cleaved from a large precursor that is processed by convertase and carboxypeptidase. Neuropeptide Y is strongly implicated in the regulation of feeding in mammals and is highly effective in stimulating GH in the goldfish (Peng, Trudeau, & Peter, 1993). However, NPY is also implicated in the regulation of GTH release (Kah et al., 1989; Breton, Mikolajczyk, Popek, Bieniarz, & Epler, 1991;

Peng et al., 1993b) and could be one of the factors establishing crosstalks between growth, feeding, and reproductive axes. Many fish species reduce food consumption during the reproductive period. Expression of NPY in the hypophysiotropic POA increases during fasting (Silverstein, Breininger, Baskin, & Plisetskaya, 1998), and stimulatory effects of NPY on LH secretion are more pronounced in fasted animals (Cerda´-Reverter, Sorbera, Carrillo, & Zanuy, 1999). In goldfish (Kah et al., 1989), NPY terminals have been reported in the pituitary, and NPY has been shown to be directly involved in the release of LH at the level of the pituitary. Neuropeptide Y is also able to release GnRH from GnRH terminals in pituitary or POA slices (Peng et al., 1993a; Trudeau, 1997). Effects of NPYon reproductive functions are modulated by sex steroids (Kah et al., 1989; Breton et al., 1991; Peng et al., 1993c). In the goldfish, pretreatment with testosterone (T) or 17b-estradiol (E2) induced a two- to threefold increase in NPY mRNA levels in the telencephalon/ POA, but not in the opticetectum/thalamus. In-situ hybridization using brains taken from T-implanted fish demonstrated that the site of steroid action is the POA (Peng, Gallin, Peter, Blomqvist, & Larhammar, 1994; Trudeau, 1997).

4.2. g-aminobutyric Acid (GABA) In the brain of all vertebrates, GABA generally is an inhibitory neurotransmitter that is extremely abundant, notably in the hypothalamus. Following the discovery of a heavy GABA innervation in the goldfish (Kah et al., 1992) (Figure 2.1 (ced)), GABA effects have been explored mainly in goldfish (Popesku et al., 2008), and to a lesser extent in rainbow trout (Oncorhynchus mykiss) (Manãnos et al., 1999). g-aminobutyric acid has been shown to stimulate LH secretion in goldfish through stimulation of GnRH release (Kah et al., 1992; Sloley et al., 1992) and inhibition of DA release (Trudeau, 1997; Trudeau et al., 2000). Intraperitoneal injection of GABA causes an increase in serum GTH in regressed

Chapter | 2


or early-maturing fish, but not in late-maturing females (Kah et al., 1992). Moreover, injection of g-vinyl-GABA (GVG), a GABA transaminase inhibitor, significantly increased GABA content in the hypothalamus and pituitary, and caused a dose-dependent increase of GTH levels (Sloley et al., 1992). The stimulatory effect of GABA on GTH release was abolished in E2-treated females, but was still observed in T-implanted fish. Moreover, E2, but not T, caused a decrease of GABA concentration within the telencephalon (Kah et al., 1992; Trudeau, 1997). In addition to modulating GABAstimulated LH release, the gonadal steroids also affect GABA synthesis in both the brain and pituitary; e.g., T and progesterone (P4) decrease, and E2 increases, pituitary GABA synthesis rates in sexually regressed goldfish (Trudeau, 1997). In rainbow trout, GABA has a stimulatory action on FSH and LH secretion. This effect is more important in females and is observed only in sexually regressed animals (Trudeau, 1997). In the rainbow trout, the stimulatory action of GABA may be exerted, at least in part, directly onto the gonadotropes, which receive a heavy innervation by glutamate decarboxylase-positive fibers (Manãnos et al., 1999). gaminobutyric acid stimulates both basal and GnRH-induced FSH and LH secretion from dispersed pituitary cells, and this effect is enhanced following E2 or T treatment (Manãnos et al., 1999). In agreement with an interaction between GABA and estrogens, estrogen receptor-a (ERa) is found in many glutamic acid decarboxylase (GAD)-expressing neurons of the hypothalamus (I. Anglade & O. Kah, unpublished results).

4.3. Gonadotropin-inhibiting Hormone (GnIH) Gonadotropin-inhibiting hormone (GnIH) is a member of the LPXRF-amide family of peptides. This dodecapeptide was recently identified in birds, in which its effects are best documented (Tsutsui et al., 2007; see also Volume 4, Chapter 1), but also in amphibians, mammals, and even invertebrates (Kriegsfeld et al., 2006; Tsutsui & Ukena, 2006; Tsutsui et al., 2007). The main role of this peptide, expressed in hypothalamic and septal neurons in birds, is to inhibit the synthesis and release of GTHs by direct actions at the pituitary level (Yin, Ukena, Ubuka, & Tsutsui, 2005; Tsutsui & Ukena, 2006; Tsutsui et al., 2007). In addition, GnIH seems to exert neuromodulatory actions on GnRH (Bentley et al., 2006; Bentley et al., 2008) and inhibits steroidogenesis and development in avian gonads, indicating that this neuropeptide could act at different levels in the reproductive axis (Tsutsui et al., 2007). The pineal hormone melatonin (MEL) may be a key factor controlling GnIH function. In the quail, GnIH neurons express MEL receptors (MEL-R) and MEL treatment stimulates expression of GnIH mRNA and mature GnIH peptide (Tsutsui

23

et al., 2007). The existence of GnIH seems not to be restricted to birds, as a peptide from the LPXRF-amide family recently has been described in fishes (Sawada et al., 2002; Osugi, Ukena, Sower, Kawauchi, & Tsutsui, 2006; Tsutsui et al., 2007). In the goldfish, mass spectrometric analyses have revealed that a tridecapeptide is an endogenous ligand (SGTGLSATLPQRF-NH2) derived from the GnIH precursor in the brain. Southern blotting analysis of reverse-transcriptase-mediated PCR products has demonstrated a specific expression of the goldfish peptide gene in the diencephalon. Immunoreactive cell bodies were found in the nucleus posterioris periventricularis and the nervus terminalis, and ir fibers were distributed in several brain regions including the nucleus lateralis tuberis pars posterioris and pituitary. Thus, the goldfish hypothalamus expresses a novel neuropeptide containing the C-terminal -LPQRF-NH2 sequence, which may possess multiple regulatory functions and act, at least partly, on the pituitary to regulate pituitary hormone release. However, at the moment it is not clear whether GnIH acts as a releasing or inhibiting factor in teleost fishes.

5. DOPAMINE (DA), A BRAIN INHIBITOR OF REPRODUCTION 5.1. Dual Brain Control of Pituitary Gonadotropins (GTHs) in Teleosts In mammals, as in most other vertebrates, GnRH neurons ensure the direct neuroendocrine control of pituitary GTHs: LH and FSH. Accordingly, modulation of GnRH neuron activity, such as their activation at puberty or at the time of seasonal reproduction, is responsible for the regulation of reproductive function. Such a single positive neuroendocrine control is not a general feature among pituitary hormones, which can be submitted to a balanced control by both stimulatory and inhibitory neurohormones. For instance, pituitary GH is under the control of two neuropeptides, GH-releasing hormone (GHRH) and somatostatin (SS), which stimulate and inhibit GH production, respectively. Similarly, TSH is stimulated by thyrotropin-releasing hormone (TRH) and inhibited by SS. Most hypothalamic neurohormones involved in the control of pituitary hormones are neuropeptides, with the notable exception of a catecholamine, DA. Dopamine is a brain neuromediator modulating various brain functions including integration of sensory cues, locomotion, emotion, cognition, and the neuroendocrine control of the pituitary. Dopamine is thus the main regulator of pituitary PRL secretion in mammals as in teleosts, by directly inhibiting lactotropes via DA D2 receptors. Additionally, there is considerable evidence that DA is a key neurohormone for the inhibitory control of gonadotropes in teleosts.

24

Neuroendocrine control of GTH secretion in teleosts may be regulated by both GnRH stimulation, as in mammals and other vertebrates, and by DA inhibition (Peter et al., 1986; Dufour et al., 2005; Popesku et al., 2008; Dufour, Sebert, Weltzien, Rousseau, & Pasqualini, 2009). The involvement of DA in the neuroendocrine control of reproduction in teleosts provides an additional brain pathway for the integration of external and internal cues and their transmission to the HPG axis (Figure 2.4). This also may have contributed to the large diversity and plasticity of teleost reproductive cycles.

5.2. Diversity of Dopamine (DA) Roles in the Control of Teleost Reproduction Teleosts encompass more than 28 000 species displaying remarkable variety of morphology, physiology, life history, and ecology. Comparative studies are highlighting conserved or divergent regulatory processes, in particular concerning the role of DA in the control of reproduction. In adult goldfish, DA negatively interacts with GnRH in the control and timing of the last steps of gametogenesis; i.e. oocyte maturation and ovulation in females, and spermiation in males. This role of DA, first evidenced in goldfish (Chang, Yu, Wong, & Peter, 1990) was confirmed in various adult teleosts, including other cyprinids, and representative species from other teleost groups such as silurids (African catfish (Clarias gariepinus)), salmonids (rainbow trout), and percomorphs (tilapia (Oreochromis spp.), grey mullet (Mugil cephalus)). This discovery led to the use of a combination of GnRH agonists and DA D2 receptor antagonist, in general pimozide, to induce an LH ovulatory peak and spawning of mature fishes for aquaculture. In contrast, the inhibitory role of DA is absent in an atherinomorph, the Atlantic croaker, and in a percomorph, the seabream (Sparus aurata). This reveals important species-specific variations in the involvement of DA in the control of the final steps of gametogenesis (for review see Dufour et al., 2005; 2009). The possible role of DA in the control of earlier steps of gametogenesis was first investigated in the European eel, a representative species of an ancient group of teleosts (Elopomorphs), which displays a unique lifecycle with a prepubertal blockade of sexual maturation before the reproductive oceanic migration. The blockade of eel sexual maturation was shown to result from a deficient production of GnRH, as in other prepubertal vertebrates, and also from a drastic inhibitory effect of DA, counteracting the effect of GnRH (Dufour et al., 1988; Vidal et al., 2004; Dufour et al., 2005; Se´bert, Weltzien, Moisan, Pasqualini, & Dufour, 2008; Dufour et al., 2009). An inhibitory role of DA in the control of puberty was recently evidenced in another teleost, the grey mullet (Aizen, Meiri, Tzchori, LevaviSivan, & Rosenfeld, 2005; Nocillado, Levavi-Sivan,


Carrick, & Elizur, 2007). In contrast, DA was reported to play no role in the control of puberty in two other percomorphs, the striped bass (Morone saxatilis) and red seabream (Pagrus major) (Holland, Hassin, & Zohar, 1998; Kumakura, Okuzawa, Gen, & Kagawa, 2003). As for the role of DA in adult teleosts, these data indicate variations among teleost species in the involvement of DA in the neuroendocrine control of puberty. Teleost species exhibit a wide diversity of lifecycles, including various examples of natural sex reversal. In the case of coral reef wrasse species, an inhibitory role of DA in the control of sex reversal has been indicated. In the protogynous saddleback wrasse (Thalassoma duperrey), experimental studies have demonstrated that DA inhibits the onset of natural sex reversal triggered by social factors and visual cues (see Chapter 8, this volume). In the protogynous bluehead wrasse (Thalassoma bifasciatum), sex reversal could be induced by administration of LH as well as by a combination of GnRH and DA D2 receptor antagonist (for review see Dufour et al., 2005). These neuroendocrine investigations reveal that DA and GnRH may interact in the neuroendocrine control of sex reversal and the mediation of species-specific triggering cues.

5.3. Evolutionary Origin of Dopamine (DA) Inhibitory Control of Reproduction in Teleosts Neuroanatomical and experimental investigations performed in goldfish have demonstrated that DA neurons originate from a specific region of the POA (nucleus preopticus anteroventralis (NPOav)) and project directly to gonadotropes in the PPD and inhibit reproduction (Kah, Chambolle, Thibault, & Geffard, 1984; Kah et al., 1984; 1987). This NPOavhypophysial DA pathway has also been characterized in some other teleost species, such as the rainbow trout (Figure 2.5(a)) (Linard et al., 1996; Ve´tillard, Benanni, Saligaut, Jego, & Bailhache, 2002), the European eel (Vidal et al., 2004; Weltzien et al., 2006), and the zebrafish (Figure 2.5(b)). In rainbow trout (Linard et al., 1996) and zebrafish (Figure 2.5(c)), such neurons express esr1 (ERa). The finding of a similar NPOavehypophysial DA pathway in various teleosts, including the European eel (A. Anguilla), a representative species of an ancient group (Elopomorph), suggests that the differentiation and recruitment of NPOav DA neurons for the direct inhibitory control of pituitary GTHs may have emerged in an ancestral teleost and been largely conserved during teleost evolution, with some variations. In rainbow trout (Linard et al., 1996) and zebrafish (Figure 2.5(c)), such neurons were shown to strongly express ERa. The above-mentioned studies highlight that the recruitment of this DA inhibitory pathway for the control of various key reproductive steps,

Chapter | 2


25

FIGURE 2.5 (a) Distribution of tyrosine hydroxylase (TH) neurons in the forebrain of the teleost fishes (Kah et al., unpublished). Although many TH-positive neurons are observed, only part of them express dopamine and very few project to the pituitary. This is notably the case of the neurons located in the nucleus preopticus anteroventralis (NPOav in black), as shown in goldfish (Kah et al., 1984a, Kah et al., 1986; Kah et al., 1987). (b) shows immunocytochemical localization of TH neurons in the NPOav of the zebrafish. This small population of DA neurons is located in the rostral preoptic area in a ventral position, above the optic chiasma (OC). (c) As also shown in trout (Linard et al., 1996), such neurons in zebrafish strongly express esr1 (Adrio & Kah, unpublished). (d,e) show TH messengers (arrowheads) detected using radioactive in-situ hybridization and esr1 mRNAs (small arrows) revealed using a dogoxygeninlabelled probe. All TH mRNA-expressing neurons also express esr1. Hc: caudal hypothalamus; Hv: ventral hypothalamus; NAPv: nucleus anterioris periventricularis; NPPv: nucleus posterioris periventricularis OC: Optic chiasma, Olf B: olfactory bulbs; Pit: pituitary; PT: posterior tuberculum; PTec: pretectum; PVO: paraventricular organ; Tec: optic tectum; VC: valvulae of the cerebellum; Vc: central part of the subpallium; Vd: dorsal part of the subpallium. Bar = 100mm

such as onset of puberty, induction of spawning, or even natural sex reversal, may have contributed to the large diversity and plasticity of teleost lifecycles. Pharmacological studies, using various DA agonists and antagonists, performed in vivo, as well as in vitro on pituitary cells, have demonstrated that DA receptors pharmacologically related to mammalian type 2 receptors (DA D2R) mediate the inhibitory action of DA on pituitary GTH secretion in goldfish (Chang et al., 1990). This is seen in all teleost species investigated in relation to the roles of DA during the final stages of gametogenesis in adult fish, in the blockade of puberty in juvenile fish, and in natural sex reversal in some protogynous species. Molecular characterization of DA receptors and localization of their expression confirm the involvement of specific DA D2Rs in the direct inhibition by DA of teleost pituitary gonadotropes (for review see Pasqualini et al., 2009). These data infer that the emergence of DA neuroendocrine control of GTH secretion in an ancestral teleost has

involved both the recruitment of a specific NPOave hypophysial DA pathway and the expression of DA D2Rs by gonadotropes. Interestingly, an investigation in a chondrostean fish, the white sturgeon (A. transmontanus) indicated an inhibitory effect of DA on GnRH-stimulated GTH secretion, exerted via D2Rs, similar to the situation in teleosts (Pavlick & Moberg, 1997). This suggests that DA neuroendocrine control of GTH secretion may be more ancient than the teleost lineage and that it emerged in an actinopterygian ancestor common to teleost and chondrostean lineages. Comparative studies could address the possible homology between brain nuclei containing DA neurons involved in the inhibitory control of reproduction in teleost and chondrostean fishes. This would indicate whether DA inhibitory control of reproduction in these groups may have resulted from a common ancestral regulatory mechanism or may have been acquired independently in the two lineages.

26

5.4. Modulation of Dopamine (DA) Inhibitory System by Internal Factors Variations in the DA control of reproduction, according to teleost species or reproductive stage, may result from variations in the activity of the NPOav neurons and/or in the expression of DA D2Rs by gonadotropes (Vacher, Mananos, Breton, Marmignon, & Saligaut, 2000; Vacher, Ferriere, Marmignon, Pellegrini, & Saligaut, 2002). As a full component of the HPG axis, the DA system is a potential target for sex steroid feedbacks. Estradiol treatment increases dopaminergic activity in adult goldfish (Dulka, Sloley, Stacey, & Peter, 1992), stinging catfish (Heteropneustes fossilis) (Senthilkumaran & Joy, 1995), and rainbow trout (Linard, Bennani, & Saligaut, 1995; Saligaut et al., 1998; Vacher et al., 2002), as measured by brain DA turnover and expression or enzymatic activity of tyrosine hydroxylase (TH). Further, DA neurons of the NPOav have been shown to specifically express ERs in the rainbow trout, indicating a direct effect of E2 on these neurons (Linard et al., 1996). When tested, T produced a similar effect to E2 (Trudeau, Sloley, Wong, & Peter, 1993), which likely results from local aromatization of T into E2 (Menuet et al., 2005; Pellegrini et al., 2005). As compared to mammals, most teleosts present a remarkably high brain aromatase (P450aro) activity, which would be related to duplication of the P450aro gene in teleosts (see Section 7) and specific expression of one of these variants in the brain. In adult teleosts, the positive feedback of sex steroids on DA neurons is interpreted as a stimulation of the dopaminergic inhibitory tone that becomes maximal at the end of gametogenesis and should drop under the control of internal (e.g., preovulatory decrease in E2) and/or environmental (temperature, pheromones etc.) cues to allow the induction of ovulation or spermiation. In contrast, an androgen-specific stimulatory regulation of DAergic neurons has been demonstrated in the prepubertal European eel. Testosterone and dihydrotestosterone (DHT) (a nonaromatizable androgen) had a similar stimulatory effect on TH expression in some brain regions including the NPOav, while E2 had no effect (Weltzien et al., 2006). This androgen-specific regulation in the eel may be related to the low brain P450aro enzymatic activity and the lack of a duplicated P450aro variant in this species, in contrast to other teleosts (Jeng, Dufour, & Chang, 2005). In male or female European eels, androgens may play a major role in the induction of morphological, physiological, and behavioral changes occurring during prepubertal metamorphosis or ‘silvering,’ which prepares them for the oceanic reproductive migration (Lokman et al., 2001; Aroua et al., 2005). The positive effect of androgens on DA neurons, associated with their various roles in the silvering process, may be interpreted as an early mechanism setting DA inhibitory tone, which prevents sexual


maturation until suitable conditions of the oceanic migration and spawning ground occur. Steroids can also exert their feedback actions on gonadotropes, by regulating the expression of DA D2Rs. Estradiol increases the expression of DA D2Rs in the tilapia pituitary (Levavi-Sivan & Avitan, 2005; Levavi-Sivan, Biran, & Fireman, 2006). Recent studies in the European eel have demonstrated the presence of two DA D2R subtypes (DA D2RA and DA D2RB), revealing a higher molecular diversity of DA D2Rs than in mammals. An androgen-specific stimulation of the expression of DA D2RB subtype, with no effect on DA D2RA, was demonstrated in some specific brain regions (Pasqualini et al., 2009). European eel DA D2R subtypes display both a differential tissue distribution and a differential regulation by sex steroids, a subfunctionalization that may have represented a significant driving force for the conservation of these duplicated genes in this species. Dopamine D2R variants also occur in zebrafish. The presence of two DA D2R subtypes in European eels should prompt further investigation in other teleosts. As for the regulation of the HPG axis, various internal factors mediating the interactions among stress, metabolism, and reproduction are likely to modulate the DA inhibitory tone. Very few studies have addressed this question in teleosts. In-situ hybridization showed that all DA neurons of the NPOav in rainbow trout express glucocorticoid receptor (GR) (Teitsma et al., 1999). Sequence analysis of the promoter of the DA drd2 gene in grey mullet revealed a putative binding site for GR (Nocillado et al., 2005). These data suggest that cortisol may control DA inhibitory tone by acting both on the activity of NPOav DA neurons and on the expression of their receptors. Future studies should investigate the impact of metabolic cues such as leptin, ghrelin, insulin, and insulin-like growth factor (IGF) on the DA system in teleosts.

5.5. Modulation of Dopamine (DA) Inhibitory System by Environmental Cues Reproduction in teleosts is closely dependent on environmental conditions for many aspects such as sex differentiation and sex reversal, onset of puberty, seasonal reproduction, induction of ovulation or spermiation, and synchronization of spawning. A large variety of environmental factors may be involved, considering the remarkable diversity of teleost species, lifecycles, and ecology. They include abiotic factorsdsuch as temperature, photoperiod, salinity, pH, oxygen, rainfall, currents, and pressuredas well as biotic factorsdsuch as nutrients, social interactions, pheromones, etc. In mammals, environmental cues are processed by the brain and ultimately integrated into the rhythmic activity of GnRH neurons. The occurrence in teleosts of the dual GnRH/DA neuroendocrine control of reproduction

Chapter | 2


provides a double brain pathway for the integration of environmental cues, and opens the way for future investigation of the impact and selectivity of various environmental factors on the teleost DA system. Among the environmental factors potentially modulating reproductive functions, photoperiod and its mediator, MEL, have been largely investigated in mammals for their role in the control of seasonal reproduction. Photoperiod is known as the major regulator of MEL secretion, but other environmental factors also have been reported to control MEL secretion, which may be especially relevant in lower vertebrates. These factors include temperature, hypoxia, exposure to magnetic fields, water pressure, and phytoplanktonic blooms. This suggests that MEL acts as a general mediator for multiple environmental cues. Acute MEL treatment reduces hypothalamic and pituitary DA metabolism in rainbow trout (Hernandez-Rauda, Miguez, Ruibal, & Aldegunde, 2000) and inhibits TH enzymatic activity in various brain regions in cycling female stinging catfish (Chaube & Joy, 2002). In mature female carp, MEL inhibited hypothalamic DA release in vivo and in vitro (Popek, Luszczek-Trojnar, Drag-Kozak, FortunaWronska, & Epler, 2005; Popek, Luszczek-Trojnar, DragKozak, Rzasa, & Epler, 2006). In contrast, in the prepubertal European eel, chronic MEL treatment enhanced the expression of TH in various brain regions including the POA, and induced a decrease in GTH expression and sex steroid levels (Se´bert et al., 2008a). This suggested reinforcement by MEL of the inhibitory DA system that blocks the HPG axis. These data suggest that MEL may exert stimulatory or inhibitory effects on the DA system according to species and physiological stage. This MELeDA control may represent an important pathway by which photoperiod and other environmental factors may modulate reproductive functions in teleosts.

6. KISS, A NEW ACTOR IN THE BRAIN’S CONTROL OF REPRODUCTION 6.1. Discovery of the Indispensable Role of KISS in the Control of Reproduction in Mammals Kisspeptins (Kps) are a family of peptides encoded by the Kiss1 gene and belonging to the superfamily of RF-amide peptides (for review see Ebling & Luckman, 2008). The Kiss1 gene and its peptides were first characterized in 1996 for their potential role in cancer as metastasis suppressors; hence, they initially were called ‘metastins.’ The reproductive role of Kps and their receptor GPCR54 (a former orphan receptor ¼ G-protein-coupled receptor 54) was unveiled when several research groups discovered that mutations in GPCR54 lead to a complete impairment of

27

reproductive function in humans and in mice (De Roux et al., 2003; Funes et al., 2003; Seminara et al., 2003). This discovery prompted an increasing number of investigations on the expression, mechanism of action, regulation, and role of Kp/GPCR54 in the control of reproduction in various mammalian models. These studies led to the demonstration that hypothalamic Kp neurons play a crucial role in the activation of the HPG axis by directly controlling the activity of GnRH neurons. Kisspeptins and their receptor GPCR54 are now considered key players and gatekeepers of various aspects of the regulation of reproductive function in mammals, such as onset of puberty, induction of ovulation, mediation of steroid feedback, and integration of metabolic and environmental cues (for reviews see Roa, Aguilar, Dieguez, Pinilla, & Tena-Sempere, 2008; Roa et al., 2009).

6.2. Investigation of the Kisspeptin (Kp) System in Teleosts The important molecular conservation of GPCR54 allowed the identification of homologous sequences in various species of vertebrates including fishes (Lee et al., 2009). After cloning of GPCR54 in tilapia (Oreochromis niloticus) and using the innovative technology of laser-capture microdissection of single digoxigenin-labeled GnRH neurons, Parhar, Ogawa, and Sakuma (2004) showed that GPCR54 was expressed by tilapia GnRH neurons. Further, GPCR54 expression was detected in each of three types of GnRH neuron present in this species, and varied between immature and mature fish (Parhar et al., 2004). This suggests that the role of Kp in the neuroendocrine control of teleost reproduction via a direct action on GnRH neurons evolved in fishes prior to the origin of teleosts and has been conserved in all vertebrates. Expression of GPCR54 changes in relation to pubertal development in the grey mullet (Nocillado et al., 2007). A similar relationship was reported in other teleost species, such as O. niloticus (Martinez-Chavez, Minghetti, & Migaud, 2008), the fathead minnow (Pimephales promelas) (Filby, Van Aerle, Duitman, & Tyler, 2008), and zebrafish, (Biran, Ben-Dor, & Levavi-Sivan, 2008). Although a single form of GPCR54 occurs in mammals, two GPCR54 receptor genes (kiss1ra, kiss1rb) are present in zebrafish (Biran et al., 2008) and two isoforms are generated by alternative splicing of a single gene in the Senegalese sole (Solea senegalensis) (Mechaly, Vinas, & Piferrer, 2009). In zebrafish, kiss1ra and kiss1rb show a differential tissue distribution, with both types being expressed in the brain, but only kiss1ra in the gonads and only kiss1rb in the pituitary and various peripheral tissues. kiss1ra and kiss1rb also differ in their transduction activity when recombinant receptors are expressed in COS cells (Cv-1 (simian) in Origin and carrying Sv40 genetic material) (Biran et al., 2008). In the Senegalese sole, a single kiss1-receptor gene generates two

28

messenger RNA isoforms after alternative splicing due to the retention of an intronic sequence (Mechaly et al., 2009). The two mRNA isoforms show a differential tissue expression. The sequence of the longer mRNA includes stop-codons, suggesting that it may lead to a truncated protein. These discoveries may promote reinvestigation of possible similar processes in other teleosts and even mammals. The cloning of Kps in fish was a more difficult task due to large sequence divergence between Kp precursors even among mammalian species. In teleosts, Kp was first identified in silico in model species in which genome sequences are available, such as zebrafish (Biran et al., 2008; Van Aerle, Kille, Lange, & Tyler, 2008) and medaka (Kanda et al., 2008). Even though the precursor sequence is highly variable, the minimal 10-amino-acid functional sequence of Kps appears highly conserved among vertebrates (Elizur, 2008; Zohar et al., 2010). Further investigation revealed the presence of a second gene coding for Kp in the sea bass (Figure 2.6), as well as in zebrafish and medaka (Felip et al., 2009; Kitahashi, Ogawa, & Parhar, 2009; Lee et al., 2009). The products of kiss1 (Kiss1) and kiss2 (Kiss2) were shown to be differentially expressed in brain regions in zebrafish as well as medaka (Kanda et al., 2008). In-vivo experimental studies showed that injection of Kiss2 was more effective than Kiss1 in stimulating LH and FSH release in sea bass (Felip et al., 2009), and in stimulating the expression of GTH subunits in zebrafish (Biran et al., 2008). In contrast, no effect of Kiss1/ Kiss2 injection was observed on GnRH expression in zebrafish in the same experiment, which may suggest that Kps would affect GnRH release more than GnRH expression in teleosts as in mammals. Further studies are necessary to characterize the mechanism of action of kiss in teleosts. Only a few studies have investigated the potential roles of internal and environmental factors in the control of Kp system in teleosts. As for the other brain and pituitary components of the HPG axis, the Kp system is an obvious target for feedback of sex steroids. In medaka, ovariectomy and E2 replacement experiments reveal that ovarian estrogens are positively regulated by Kp expression in a specific hypothalamic nucleus, the nucleus ventral tuberis (NVT), with no effect on Kp neurons from another hypothalamic nucleus, the nucleus posterioris periventricularis (Kanda et al., 2008). Accordingly, variations in the expression of kiss in the NVT were observed at different stages of the reproductive cycle (Kanda et al., 2008). Concerning environmental factors, light regime was shown to affect the expression of GPCR54 in Nile tilapia (Martinez-Chavez et al., 2008), which suggests that the Kp system may be one pathway for photoperiod/MEL modulation of reproduction in teleosts, as indicated in seasonal mammals such as the Syrian hamster (Revel et al., 2006; for review see Elizur, 2008). Future studies should aim at investigating the effects and mechanisms of action of various internal (including


metabolic clues) and environmental factors on the Kp system in teleosts.

6.3. Origin and Evolution of the Kisspeptin (Kp) System Although further research is needed, we may assume that there is a functional Kp system in teleosts. At the moment, it is unclear to what extents the functions of Kp in fishes are similar to those known in mammals. Indeed, while there is evidence for the presence of kiss genes in the common ancestor of the actinopterygian and sarcopterygian lineages, there seems to be quite a lot of variability in the number of kiss genes in vertebrates. Figure 2.7 shows the phylogenetic analysis and genomic organization of vertebrate kiss1 and kiss2 genes. The current information indicates that two kiss genes were present in basal vertebrates such as the sea lamprey (Petromyzon marinus) (agnathan). Two genes are present in most modern fishes examined and two or three kiss genes are found in the amphibians, Xenopus laevis and Xenopus tropicalis, respectively. There are two in the prototherian mammal platypus (Ornithorhynchus anatinus) (Felip et al., 2009; Lee et al., 2009), but in contrast only one kiss gene has been found in modern reptiles such as the grass lizard (Takydromus tachydromoides) and eutherian mammals (Lee et al., 2009). Similarly, it appears that some modern fishes may have lost one copy, as only the kiss-2 gene has been found in the fugu genome (Figures 2.6 and 2.7). So far, no kiss gene has been reported in birds. Analysis of genome sequences has revealed the presence of GPCR54-like gene sequences in representative species of groups that emerged even before the vertebrate lineage. This is the case for GPCR54-like sequences detected in an echinoderm, the sea urchin (Strongylocentrotus purpuratus) (for review see Biran et al., 2008). At the moment, no Kp-like sequence has been found in these species, which may be due to large divergences among kiss gene sequences. If the presence of a Kp/GPCR54 system is verified in these species, the next challenge would be to investigate whether there is a functional link between Kp/GPCR54 and GnRH systems in these nonvertebrate species. Gonadotropin-releasing hormones and/or GnRH receptors have been characterized in amphioxus, ascidians, and mollusks (for reviews see Kah et al., 2007; Zohar et al., 2010). The presence of a GnRH-like peptide with a potential role in reproduction also has been suggested in cnidarians, one of the most ancient groups of metazoans (Twan et al., 2006). As discussed in Section 2, these data suggest an ancient evolutionary origin of the GnRH system, much before the emergence of the pituitary gland and GTHs. Therefore, it will be fascinating to retrace the evolutionary origin of the kiss system and the emergence of the functional link between Kp and GnRH systems in metazoa.

Chapter | 2

A

29


Sea bass KiSS-1 1 1 100 28 199 61 298 94 396 495

GGCACGAGCTCTCTCTCTCTCTCTCTCTCTCTCTCTCTCTCTCTCTCTCTCTCTCTCTCTCTCCTCAGTCAGCACTCGCCTTCAGCACACCGAGGAATT M P R L I V A L M I A A L S T E I Y N T S M I S S Y H TCAGCAGGTCTGTCACGATGCCCCGACTCATTGTCGCTCTGATGATAGCTGCTTTGTCAACAGAGATCTACAACACCAGCATGATATCCAGCTACCACA S K ▼D Q V I L K A L R D L S H A S I L A S A K N S G N L P A D K V GTAAAGATCAGGTAATACTCAAAGCCCTGAGAGATTTAAGCCATGCATCAATACTGGCATCAGCAAAGAATTCCGGGAATTTACCTGCTGATAAGGTCC H S A D G K F P R S E W L I S K L V L P Q T I K K R Q D V S S Y N ATTCAGCTGATGGAAAGTTTCCCAGGTCAGAATGGTTGATCTCAAAGCTGGTCCTCCCTCAGACTATCAAGAAACGTCAAGATGTGTCCTCATACAACC L N S F G L R Y G K * TCAACTCCTTTGGTCTACGTTACGGAAAGTGACACAAGACCTGATGTCTGTTGTTATTCTGTGTTAGAGCTGATATTTTTATATATTTTCTTTGTACAT TCAGAGTGGGAAAAGAAAAGTGTAATGTCACTGTTGAAAGCTCAATAAAAATGTTGGTTAAAGATT

Sea bass KiSS-2 1 1 25 100 58 199 91 298 396 495 595 694

B

M R L V A L V V V C G L I L G Q D G G S V G A A GGCACGAGGAGACACACACACAGAGGATGAGGCTCGTGGCTCTGGTCGTGGTGTGCGGGCTGATCCTGGGTCAGGATGGAGGGAGCGTGGGAGCAGCTC L P E L D S A Q R T G A T ▼G S L L S A L R R R T A G E F F G E D S TGCCGGAACTCGACTCTGCACAGAGGACAGGAGCAACAGGTTCACTCCTCTCTGCGCTCAGGAGGAGGACTGCAGGAGAGTTTTTTGGGGAGGATTCCA S P C F S L R E N E E Q R Q L L C N D R R S K F N F N P F G L R F GCCCGTGTTTCTCTCTGAGAGAGAACGAGGAGCAGCGGCAGCTCCTGTGCAACGACCGCAGGAGTAAATTCAACTTCAACCCGTTCGGCCTCCGCTTCG G K R Y I Y R R A L K R A R T N R F S P L F L F S R E L E V P T & GGAAGCGCTACATTTACAGAAGGGCCCTTAAAAGAGCCAGGACGAACAGGTTCTCGCCCCTTTTTCTCTTCTCACGAGAACTGGAGGTGCCTACCTGAT GCCTACTGATGTGTCTTCCTCTGAGGACTTGTGTCCCATTGGTGAAAAGTCAACATGTGACAAGTCTTTCCATGTGTTTAAAATAGGCCTTTTACTTCA GATAAAAGGTTTCGGTGAATTAAACTTTTTGTACCTACCTTTACTGTGACATGATTTGAACATGGAAGAATACGAGAGAAAAACACACTTACAACAGTG TAAACAGAATGCTTTAGACAATATCATTTTGTAAAATGTTTGCTTTTCTCATCTCAGTTTTTAAGAAATCTAAATGAAACAGTCCAATGGACTTTTATG ATGATGGGAAAGGAGTAGTTCATTTCACAATAAAAGCACTGTATGGTTAAT

KiSS-1 rat mouse human opossum platypus Xenopus seabass medaka zebrafish lamprey

MISLASWQ-LLLLLCVASFGEPLAKMAP-VVNPEPT---GQQSGPQELVNAWQKGPRYAESKP-GAAGLRARR-TSPCPPVEN-PTGHQ MISMASWQ-LLLLLCVATYGEPLAK-----VKPGST---GQQSGPQELVNAWEKESRYAESKP-GSAGLRARR-SSPCPPVEG-PAGRQ MNSLVSWQ-LLLFLCATHFGEPLEKVAS-VGNSRPT---GQQLESLGLLAPGEQSLPCTERKP-AATARLSRRGTSLSPPPES-SGSRQ MRSSVYWQ-LLLLLSVSPFGETSDKFAP-VENPGRT---GQPGRLLAHLIPWEGRPQCLEK---PEQTGQTQRLAMLCPSDE--ASDPL MSSLTSLL-LLLFLCAFPFGETVEKIHPPFKTPDFQ---RTRGQWERTPGQWERTPPCLDKKPGPEAAEQTARLALLCPPEE--SPGQV ---------------------------------------DELYSQVPGKSQWLGSLLCPEKVPTTRRAEQMPVLSLLCRRKKSLSTGHP MPRLIVAL-MIAALSTEIYNTSMISS---YHSK--------DQVILKALRDLS-----------HASILASAKNSGNLPADKVHSADGK MAAPLIVAVIMWAVLAQVWTAHHRHQST-IHTE--------DNALLKMLRNFN-----------YLS--SSMKEWP--KSDR--SSDGG ---MMLLTVILMLSVARVHTNPSGHFQY-YLE--------DETPEETSLRVLRG---------------TDTRPTDGSPPSK--LSALF MRGLTVVTFLFLVLCCDSFGKVVSFYG--FKESTKSGGGQLPGDVTDILREITS----------LLEGTDGIVAFYDFPGSGGSVDRAF

rat mouse human opossum platypus Xenopus seabass medaka zebrafish lamprey

RP-PCATRSRLIPAPRGSVLVQREKDMSAYNWNSFGLRYGRRQVAR------AARG---------RP-LCASRSRLIPAPRGAVLVQREKDLSTYNWNSFGLRYGRRQAAR------AARG---------QPGLSAPHSRQIPAPQGAVLVQREKDLPNYNWNSFGLRFGKREAAPGNHGRSAGRGWGAGAGQ--WPGLCPTRSRLITAPQGALLVEREKDMSTYNWNSFGLRYGKRQTNRG------------------WRGLCPTQSQLVTGPQGGMLVEQEKDLSAYNWNSFGLRYGKRHAGVLK---ARLKIW--------WSTDSLLPSRSISAPEGEFLVQREKDLSTYNWNSFGLRYGKRGSGSEN---SKTKVW--------FPRSEWLISKLVL-P---QTIKKRQDVSSYNLNSFGLRYGK------------------------TPMVGCWMVKALH-P---VAIKKRQDLSSYNLNSFGLRYGK------------------------SMGAGPQKNTWWWSPE-SPYTKRRQNVAYYNLNSFGLRYGKREQDMLTRLKQKSPVK--------MSPLHFYPMLRARMRSLPASDADEKKGSTYNWNSFGLRFGKRELNFMNISKILIIFTKRQ-----Kisspeptin- 10

KiSS-2

81 77 82 79 83 50 66 63 60 77

130 126 145 126 137 104 103 100 116 137

platypus lizard Xenopus zebrafish seabass stickleback Fugu Tetraodon medaka lamprey

--MR--ILPFLVMGLIYQSGSLGNPLLGERKAVKRLEFTDVPAVHAEAK---------RGEQPAGQEAAAGPGLCYLVQGSRVQ---------MFRLLLLSFFVVILSPNGTFGKPVYGDFR--SILQVAFSDAADPANDLQAKRNSYLNTRESEVLDSEDPSSLCYFIQESETESQ-------MLLLLLLTLVISQHAVGGTMFR---GDEEGLELEEIGGPETSYPEGDPREKSESYELIPSADTLSWPGRSNICYFIREGRLESQ-------MN-TRALILFMSAMVSQSTAMRAILTDMDTPEPMPDPKPRFLS------------MERRQFEEPSASDDASLCFFIQEKDETSQ-------MR--LVALVVVCGLILGQDGGSVGAALPELDSAQRTGATGSLLSA----------LRRRTAGEF-FGEDSSPCFSLRENEEQRQ----------------------------------------------GSVLST----------LTRRTIGAF-LADDANPCFSLRENEEQRR-------MR--VLVLLLV--LAVAPDRGG------AHATMQVTGGSGSVQ------------LRRGTAGQLQLLQESNPCLTFRDNEDQ---------MRLWVLVLLLVRALTVAQDRGAT-----HHATVQGAGGPGSVR------------LRKGTVGEL-LLEESNPCVALRDNQDQ---------MT--RAVVLVLCALIAAQDGGRAAAGLAARDSGRGTHATGVLW------------ILRRSEDDS-AAGGAGLCSSLREDDEQ-------MTPACSLAALLAVCVFGGGAVAARTDRYGASPDSNHARRARSSEEIVTGDLRASPLRLFGAVCRHAAETPRLLRLRALRGGHDLDAGLTDGE

71 82 81 71 71 34 60 64 67 92

platypus lizard Xenopus zebrafish seabass stickleback Fugu Tetraodon medaka lamprey

-ISCRLRFTRGQFNFNPFGLRFGKR------DPGKPAPLPARAPGLPAARNSEPRGWIQCGAARGGRC------------------------132 -ISCRLRFTRSKFNFNPFGLRFGKRQGDTLADDGKLGSQGSRKILQALLKPRLDQTHSQCGENWGDTC------------------------149 -LSCHLRFTRSKFNFNPFGLRFGKRARGDANGEG-LAPLVPRRLLPFLLK----LKDKRCSESVGESC------------------------143 -ISCKHRLARSKFNYNPFGLRFGKR------------NEATTSDSDRLKHKHLLPMMLYLRKQLETS-------------------------125 -LLCND--RRSKFNFNPFGLRFGKR-------------YIYRRALKRARTNRFSPLFLFSRELEVPT-------------------------122 -LLCND--RRNQFNFNPFGLRFGKR-------------YIYRRAVKRARTNTLSPLSLRSSLFLR---------------------------83 -LLCN----RSKFNLNPFGLRFGKR-------------FIYRRAMKQARTHTRSPVSQEVPT------------------------------104 -LLCN----RSRFNLNPFGLRFGKR-------------LVYRRAMKLARTRALPPVSQEVPT------------------------------108 -LLCAD--RRSKFNYNPFGLRFGKR-------------APPPRGAHRARAMKLPLMSLFQEVPT----------------------------115 ALPRSAEQDVTEFNYNPFGLRFGRR------SGAQSSTAATRSRAEAATSRRWTSRFSRLPAVQNPEEFSALILMISAVQCQFRNVSHVFVT178 Kisspeptin- 10

FIGURE 2.6 (A) shows the sea bass KiSS-1 and KiSS-2 oligonucleotides and protein sequences. (B) shows alignments of sequences of vertebrate KiSS1 and KiSS-2 genes. The residues conserved in all species are shown in black. Residues conserved in 70% and 50% of the species are represented in dark grey and light grey, respectively. The region corresponding to kisspeptin-10 is indicated. Reproduced from Felip et al., (2009), with permission from Elsevier.

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FIGURE 2.7 (a) Unrooted neighbor-joining bootstrap consensus tree of kiss1 and kiss2 genes in vertebrates. An alignment of amino acid sequences corresponding to exon 2 was used. Ambiguously aligned sequences and gaps were taken out before analysis. (b) Synteny of KiSS-1-containing regions in vertebrate genomes. In the mouse and rat, the proximity between KiSS-1 and GOLT1A (number 7) indicates that the sequence of KiSS-1 was found within GOLT1A. Genes shown in boxes in human chromosome 1 (Chr1) are located in other chromosomes in that species. (c) Synteny of KiSS-2-containing regions in vertebrate genomes. The genes boxed in the mouse and rat are located in the indicated chromosome. Dr, Danio rerio; Fr, Fugu rubripes; Ga, Gasterosteus aculeatus; Gg, Gallus gallus; Hs, Homo sapiens; Md, Monodelphis domestica; Mm, Mus musculus; Ol, Oryzias latipes; Rn, Rattus norvegicus; Tn, Tetraodon nigroviridis. In both schematic representations, numbers on the boxes indicate different genes. Orthologous genes are represented as squares, and those cases where paralogs were found in KiSS-1 and KiSS-2 analyzed regions are represented as ovals. A black bar between genes indicates that these are not contiguous. Dashed boxes for KiSS-1 or KiSS-2 represent absence of the gene. Numbers in brackets on the right indicate which chromosome (Ch), scaffold (S), or linkage group (LG) the cluster is found on. Reproduced from Felip et al. (2009), with permission from Elsevier. See color plate section.

6.4. The Kisspeptin (Kp) System in Fishes: Missing Link Between Growth/Metabolism and Reproduction In all organisms, there is a well-known conflict between energy targeted to growth and resources mobilized for reproduction. In oviparous species such as fishes, this

conflict is even more important, given the high energetic cost of female oogenesis and the low survival of the progeny. Thus, any individual suffering reduced energy stores or metabolic stress will exhibit retarded puberty and/ or reduced fertility. As in all vertebrates, the crosstalk between energy status and the reproductive axis is ensured through sensing of metabolic signals by neuroendocrine

Chapter | 2


circuits involving neuropeptides (such as NPY, cocaineand amphetamine-regulated transcript (CART), Kp, and GnRH), and through a number of neuropeptide hormones and metabolic cues, the nature and mechanisms of action of which are still largely unknown. In this context, the emergence of Kps, encoded by the kiss1 or kiss2 genes, and their receptors, kiss1ra and kiss1rb, as mandatory signals for normal pubertal maturation and gonadal function, has raised the possibility that the Kp/GRP54 system might also participate in coupling body energy status and reproduction. In rodents, there is now strong evidence that the KiSS-1 system is crucial to mediating metabolic information onto the centers governing reproductive function, through a putative leptineKpeGnRH pathway (Roa et al., 2009). Although fish leptin has now been characterized and produced, there is still a great need for detailed information on its effects upon the neuroendocrine circuits controlling reproduction and feeding or the expression of leptin receptors (Huising et al., 2006; Nagasaka, Okamoto, & Ushio, 2006; Murashita, Uji, Yamamoto, Ronnestad, & Kurokawa, 2008; Yacobovitz, Solomon, Gusakovsky, Levavi-Sivan, & Gertler, 2008). It is possible that, as in mammals, leptin will emerge as one of the actors ensuring the dialog between the growth/nutrition axis and the reproductive axis. It would be essential to identify the sites of expression of leptin receptor in the brain of fishes and their relationships to the neuroendocrine circuits.

7. SEX STEROIDS IN THE BRAIN OF FISHES It is clear that sex steroids exert positive and negative feedback on the brainepituitary complex of fishes to modulate the activity of the neuroendocrine circuits. However, the detailed mechanisms underlying these effects are still poorly understood given the tremendous diversity of reproductive strategies in fishes, which is likely related to their high sexual plasticity. Recent studies indicate that aromatase seems to be of crucial importance in influencing the phenotypic sex of the gonad (Guiguen, Fostier, Piferrer, & Chang, 2009) and most likely that of the brain.

7.1. Aromatase Expression and High Sexual Plasticity of the Brain in Fishes In all vertebrates, P450aro activity is present in the brain and plays a critical role in sexual differentiation (Lephart, 1996). In mammals, according to the aromatization hypothesis (Phoenix, Goy, Gerall, & Young, 1959; Thornton, Zehr, & Loose, 2009; see also Volume 5, Chapter 1), the embryonic testis, upon the influence of the sex-determining factor SRY, releases T, which is aromatized in the brain. The resulting E2 then irreversibly masculinizes certain brain regions such as the dopaminergic sexually dimorphic anteroventral periventricular nucleus of the

31

hypothalamus according to ER-dependent mechanisms, as shown in mice (Simerly, Zee, Pendleton, Lubahn, & Korach, 1997). In agreement with this hypothesis, P450aro activity in the brain of rodents is maximal during the embryonic and perinatal period (Lephart, 1996). In contrast, in fishes there is no strong genetic sex-determining system and sexual differentiation is influenced by environmental factors, including social cues (Guiguen et al., 2009; Herpin & Schartl, 2009). Many fishes can change sex within their lifespan, indicating that brain sexualization is not irreversible and can occur late in life. Probably in relation to this fact, P450aro activity increases with sexual maturity to reach extremely high levels in the brain of mature animals, either males or females (Gonzalez & Piferrer, 2003). The cyp19a1 gene, which encodes P450aro, is another example of a gene that was duplicated in fishes. A subfunctionalization process possibly followed this duplication event. Indeed, the resulting copies show clearly complementary expression patterns; the cyp19a1a gene, encoding P450aro A, is mostly expressed in the gonads and the cyp19a1b gene is mostly expressed in the brain (for reviews see Pellegrini et al., 2005; Mouriec et al., 2008; Diotel et al., 2010). The latter gene, encoding another P450aro B protein with slightly different properties from P450aro A, is highly expressed in the brain of adult mature fishes. The reason for this high expression is that the cyp19a1b gene, in many teleost species but not all, is strongly upregulated by estrogens and some aromatizable androgens (Chiang et al., 2001; Tchoudakova, Kishida, Wood, & Callard, 2001; Menuet et al., 2005; Le Page, Scholze, Kah, & Pakdel, 2006; Le Page, Menuet, Kah, & Pakdel, 2008; Mouriec et al., 2009). Given this hypersensitivity to estrogens, the cyp19a1b gene is also a potential target for xenoestrogens (Tong & Chung, 2003; Cheshenko et al., 2007; Lassiter & Linney, 2007; Cheshenko, Pakdel, Segner, Kah, & Eggen, 2008; see also Chapter 13). Interestingly, P450aro B is only expressed in radial glial cells of the brain in fishes (Forlano, Deitcher, Myers, & Bass, 2001; Menuet et al., 2003; Menuet et al., 2005; Strobl-Mazzulla et al., 2008; Diotel et al., 2010). For example, in rainbow trout (Figure 2.8(aed)), the messengers are mostly observed along the brain ventricles, corresponding to messenger accumulation within the cell bodies of the radial cells. Indeed, radial cells are characterized by a small nucleus located near the ventricle and long radial extensions terminating by end feet on the pial surface (Pinto & Gotz, 2007). In zebrafish, messengers are found within the radial processes and they also accumulate in the end feet of the radial cells (Pellegrini et al., 2005; Diotel et al., 2010). In all species studied so far using immunohistochemistry, the P450aro B protein was detected in radial glial cells, including their processes and the endfeet where the protein also accumulates. Expression is observed mainly in the

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FIGURE 2.8 Expression of cypa19a1b (aed) mRNAs and protein (eef) in the brain of fishes. (a and b) Autoradiogram of a sagittal section of the brain of an adult female rainbow trout hybridized with cypa19a1b antisense (a) and sense (b) probe. Note the very strong signal in the ventral forebrain at the level of the olfactory bulbs (ob), preoptic area (poa), hypothalamus (hyp), saccus vasculosus (sv), and pituitary gland (P). The signal is weaker in the optic tectum (Otec) and cerebellum (CC). Anterior to the right. Bar ¼ 2.5 mm. (c and d) In-situ hybridization of transverse sections of a female rainbow trout showing the very strong cyp19a1b mRNA signal (arrows) over the ventricular layer at the level of the magnocellular preoptic nucleus (NPOmc) and the mediobasal hypothalamus (MBH) and nucleus recessus lateralis (NRL). Note that the signal extends radially in radial cell processes (arrowheads). Bar ¼ 50 mm. (deh) Transverse sections in the brain of a female zebrafish showing cyp19a1b immunoreactivity in radial glial cells of the caudal preoptic region (e), thalamus (e), caudal hypothalamus (f and g) and in the optic tectum (h). Hc, caudal hypothalamus; NAT, nucleus anterioris tuberis; nh, habenular nucleus; NPPv, nucleus posterioris periventricularis; NPT, nucleus posterioris tuberis; NRP, nucleus recessus posterioris; NSC, suprachiasmatic nucleus; OT, optic tract; rl, recessus lateralis; rp, recessus posterioris; Thal, thalamic region; TS, torus semicircularis; VC, valvula of the cerebellum. (e, f, h) Bar ¼ 200 mm. (g) Bar ¼ 50 mm.

anterior brain: the POA and MBH (Figure 2.8(a, ceg)). However, P450aro B positive radial cells also are observed in the mesencephalon, notably the optic tectum (Figure 2.8 (h)), and around the fourth ventricle and rostral spinal cord. Radial glial cells represent a unique cell type acting as neuronal stem cells in the developing brain of vertebrates (Pinto & Gotz, 2007). In mammals, such cells disappear at the end of embryonic neurogenesis as they transform into mature astrocytes. In contrast, radial glial cells persist in adult fishes and have recently been shown to serve as neural progenitors (Pellegrini et al., 2007) in a way similar to what is known in the embryonic mammalian brain (Noctor et al., 2002). It thus seems that the brain of adult fishes retains some properties of the developing brain in mammals, including radial glial cells showing high neurogenic activity and high P450aro B expression (Diotel et al., 2010). These unique properties not only allow the brains of fishes to grow during their entire lifespan, but also to build sex-specific

circuits as the neurogenic activity is likely to be modulated in a regional manner by the action of sex steroids. Although this is not firmly established, such a capacity to sexualize their brains throughout life would provide fishes with an exceptional sexual plasticity permitting the development of a large variety of sexual strategies, notably all types of sex change.

7.2. Classical Positive and Negative Feedbacks In teleosts, many species are asynchronous or groupsynchronous spawners, resulting in a complex pattern of sex steroid production that makes it difficult to address the steroid feedback issue. The potential feedback effects of sex steroids are thus easier to decipher in synchronous species. In rainbow trout, e.g., one of the best-studied teleosts, three

Chapter | 2


major steroids are produced in females: E2, synthesized over the vitellogenic period; T, which shows increased production during late vitellogenesis; and the maturation-inducing hormone 17a,20b-dihydroxy-progesterone (Campbell, Fostier, Jalabert, & Truscott, 1980). These hormones are key synchronizing factors for maturation of neuroendocrine circuits, their inhibition or activation at the right moments, and the synchronization of neuroendocrine events with sexual behavior. Estradiol is synthesized by the granulosa cells of the follicles during the early vitellogenic period. A key enzyme for E2 synthesis in the gonad is P450aro A, which is a product of the cyp19a1a gene in teleosts. Estradiol acts through different mechanisms, involving well-characterized intracellular estrogen receptors (ERs) (Hawkins et al., 2000; Menuet et al., 2002), belonging to the large nuclear receptor family. Fishes also have estrogen membrane receptors that are less well characterized (Thomas et al., 2006). Three ERs have been reported in many species: Esr1 (formerly ERa)dcorresponding to esr1 of tetrapodsdand two othersders2a (formerly ERb2) and esr2b (formerly ERb1)dthought to be the result of the teleost-specific genome duplication 300 MYBP (Menuet et al., 2002; Kah, 2009). These three ERs are functional in fishes and all have strong affinity for E2 in the low nanomolar range (Menuet et al., 2002). They are expressed in many target organs, notably the gonads, brain, pituitary, and liver. The best documented expression is that of the esr1 in the rainbow trout, the only species in which it has been studied at both the protein and mRNA levels in the same species (Salbert et al., 1991; Salbert, Atteke, Bonnec, & Jego, 1993; Anglade et al., 1994; Pakdel et al., 1994; Navas et al., 1995; Linard et al., 1996; Menuet, Anglade, Flouriot, Pakdel, & Kah, 2001; Menuet, Adrio, Kah, & Pakdel, 2004). In the pituitary, Esr1 is expressed in LH cells and is involved in the expression of the b-subunit of the LHb gene (Breton & Sambroni, 1996). Although esr2a and esr2b are also expressed in the pituitary, their precise sites of expression are poorly documented. In the brain, the strongest esr1 expression is in the ventral telencephalon, the POA, and the MBH (Salbert et al., 1991; 1993; Anglade et al., 1994; Pakdel et al., 1994; Navas et al., 1995; Menuet et al., 2001; 2002; 2003; 2004). These regions also express esr2a and esr2b, as shown in a few species. In zebrafish, esr2a and esr2b have a wider distribution than esr1 (Menuet et al., 2002). They are notably present above the periventricular areas of the telencephalon and diencephalon, where they are likely implicated in the estrogenic regulation of cyp19a1b (aromatase B) expression. As observed in mammals, esr2a and esr2b expression are found often in the same cells (Kah, 2009). In rainbow trout, esr1 mRNAs also have been detected by Northern blotting in the retina and pineal, where E2 is able to modulate MEL secretion (Begay et al., 1994).

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During vitellogenesis, E2 plasma levels increase and cause vitellogenin synthesis in the liver through a well-known Esr1-dependent mechanism (Pakdel, Le Gac, Le Goff, & Valotaire, 1990; Pakdel, Feon, Le Gac, Le Menn, & Valotaire, 1991). In addition, E2 exerts simultaneous effects on LHb subunit synthesis in the pituitary (Saligaut et al., 1998) and GnRH synthesis in the brain (Montero et al., 1995; Dubois, Florijn, Zandbergen, Peute, & Goos, 1998). As stated above, E2 reinforces dopaminergic inhibition by increasing DA turnover and increasing DA D2R expression (Vacher et al., 2000; 2002). At the end of seasonal vitellogenesis, ovarian P450aro A expression is turned off, therefore favoring the production of T. A substantial part of T actions on the brain are mediated through aromatization into estrogens. This is notably the case for T effects on GnRH synthesis that are not observed with nonaromatizable androgens (Breton & Sambroni, 1996; Dubois et al., 1998). However, in the goldfish or rainbow trout, T treatment causes increased sensitivity of the gonadotropes to GnRH, an effect that is not mimicked by E2 treatment. In the goldfish, microarray expression profiling of the telencephalon identified 98 differentially expressed genes after fadrozole (an aromatase inhibitor) treatment (Popesku et al., 2008; Zhang et al., 2009a; 2009b). Some of these genes are estrogen-responsive in fishes and other vertebrates, including the rat, mouse, and human. Gene ontology analysis together with functional annotations have revealed several regulatory themes for physiological estrogen action in fish brain, which include the regulation of calcium signaling pathways and autoregulation of ER actions. Androgens in fishes can be divided into aromatizable (e.g., T, androstenedione) and nonaromatizable (e.g., DHT, 11-KT). They act through androgen receptors (ARs), of which two forms occur in some species (Takeo & Yamashita, 1999; Todo, Ikeuchi, Kobayashi, & Nagahama, 1999; Sperry & Thomas, 2000; Blazquez & Piferrer, 2005; Burmeister, Kailasanath, & Fernald, 2007) but only one in others, such as zebrafish (De Waal, Wang, Nijenhuis, Schulz, & Bogerd, 2008). According to recent studies, three events have shaped the present diversity of ARs in actinopterygians: (1) the teleost-specific genome duplication, (2) the loss of one duplicate in several lineages, and (3) the putative neofunctionalization of the same duplicate in percomorphs (Douard et al., 2008). Whereas nonaromatizable androgens most likely act through ARs, aromatizable androgens can be aromatized due to the high expression of the cyp19a1b gene in the brain of fishes (Pellegrini et al., 2007; Mouriec et al., 2009). The main brain areas showing expression of ARa and ARb are the POA and the ventral hypothalamus (Menuet et al., 1999; Harbott, Burmeister, White, Vagell, & Fernald, 2007). In the pituitary, ARa and ARb are expressed with similar patterns, but ARa mRNA is present at a higher level. These results are in total agreement with those obtained in rainbow trout. In this latter species, both ARa and ARb

34

(Takeo & Yamashita, 1999) are detected in the pallial regiondthe POA, MBH, optic tectum, and pituitaryd where ARa messengers most likely overlap with the gonadotropes. Expression of ARa in the gonadotropes is consistent with the fact that in rainbow trout T, but not E2, strongly increases LH release, indicating a clear androgenic effect at the gonadotrope (Breton & Sambroni, 1996). In contrast to most teleosts, studies in the Japanese eel (Anguilla japonica) have revealed a low brain P450aro enzymatic activity, with no evidence for the presence of the duplicated cyp19a1 gene (Jeng et al., 2005). Japanese and European eels also present the peculiarity of having high androgen plasma levels (T and 11-KT) in the female as well as in the male. Accordingly, in these species various brain functions are regulated by androgen-specific mechanisms, as exemplified by the dopaminergic system. In the European eel, E2 fails to upregulate tyrosine hydroxylase expression in the preoptic area, whereas T is successful (Weltzien et al., 2006; Pasqualini et al., 2009). Androgens are also believed to play a key role in both sexes in the induction of the prepubertal metamorphosis called silvering, preparing the eels for the spawning migration (Aroua et al., 2005). The appearance of androgen-specific regulation in this species may have emerged in relation to its unique lifecycle. Aromatizable androgens can be converted to estrogens in the brain, given the high expression of the cyp19a1b gene (Mouriec et al., 2008). However, recently it was shown in zebrafish that the brain can convert DHT into 3b-diol through the action of the enzyme 3b-hydroxysteroid dehydrogenase (3b-HSD) (Mouriec et al., 2009). The resulting 3b-diol is an androgen with potent estrogenic activity. 3b-diol binds strongly to all estrogen receptors in both fishes and mammals (Mouriec et al., 2009), providing another pathway through which T can be converted into an estrogenic compound. Thus, earlier observations employing DHT where conclusions were based on the nonaromatizability of DHT may require reinterpretation.

7.3. Expression of Other Steroidogenic Enzymes in the Brain of Fishes There is a strong correlation between the sites of expression of P450aro and that of sex steroid receptors. However, recent data also suggest that brains of adult fishes express other steroidogenic enzymes such as the cholesterol side-chaincleaving enzyme (P450scc), 17-hydroxylase 17,20 lyase (P450c17), 3b-HSD, 5a-reductase, and 17b-hydroxysteroid dehydrogenase (17b-HSD) (N. Diotel et al., unpublished). This suggests that the brain of fishes not only produces estrogens, but also a variety of other sex steroids. In addition, it is known, e.g., that ERs, ARs, and PRs are also strongly expressed in brain regions expressing aromatase and other steroidogenic enzymes. Further, some of these enzymes are expressed in a sexually dimorphic manner in


the developing brain of rainbow trout or in sex-changing black porgy (A. schlegeli) (Tomy, Wu, Huang, Dufour, & Chang, 2007). Certainly, these exciting results await confirmation and also more research before one can draw a clear picture from these data. However, the roles of these neurosteroids can no longer be neglected.

8. CONCLUSIONS AND PERSPECTIVES Fish reproductive neuroendocrinology is still a young science if one considers that it was born during the 1970s, with notably the first article on luteinizing hormonereleasing hormone (LHRH) in carp (Breton et al., 1971), the first purification of fish GTH in carp (Burzawa-Gerard, 1971), and the first brain-lesioning studies in goldfish (Peter & Gill, 1975). Today, after one generation of active researchers, the amount of information available is rather impressive. The classical techniques of physiology based on the measurement of hormones have been enriched by the molecular biology approach, allowing for rapid acquisition of sequence information for studies of gene expression and gene regulation. The sequencing of an increasing number of fish genomes now permits performance of pangenomic studies that aim to understand how gene networks are organized and how they collectively respond to environmental and physiological challenges. This allows access to a new dimension in our understanding of the functioning of organisms. The benefits of these extremely powerful approaches are yet to come and will be proportional to our mastering of this huge amount of data and our capacity to integrate all levels of information. This is the challenge of ‘integrative biology,’ the new way to approach ‘physiology.’ In addition, the richness of the fish field also comes from the multiple model species that are worked on, from basal teleosts such as the arowanas (Osteoglossidae) and the eels (Anguillidae) to the more recent fugu or flatfishes. This provides us with the unique opportunity to compare, with the same level of resolution, species separated by possibly 200 million years of evolution, and thus to gain access to the evolutionary dimension that makes biology such a fascinating science. Certainly, more research on agnathans, chondrichthyans, primitive actinopterygians, and sarcopterygians would contribute to our understanding of the evolution of these systems. Additionally, this progress has increased considerably our capacity to manage fishes in captivity, to diversify the cultured species, and to increase the overall production of farmed fishes. Compared with other sectors of aquaculture, finned-fish farming is an extremely rapidly growing sector, although there are still a number of bottlenecks that vary from species to species. The main problems include precocious sexual maturation, as is the case in salmonids; skewed sex ratios as in sea bass; and total (eel) or partial (trout) infertility in fish farming conditions. Future research

Chapter | 2


should notably aim at a better mastering of the factors triggering puberty in fishes and also at deciphering the influence of environmental factors in the control of gonadal and brain sex differentiation. Another important area for future investigations relates to the control of sexual behavior, which is often impaired in controlled environments and causes lack of spawning in some species.

ABBREVIATIONS 17,20b-DHP 17b-HSD 3b-HSD ACTH AR CART cGnRH-II DA DHT E2 ER ERa FSH GABA GAD GAP GH GHRH GKR GnIH GnRH GnRH-R GPA2 GPB5 GPCR GPCR54 GR GTH GVG HPG IGF ir Kp LH LHRH MBH ME MEL MEL-R mGnRH MYBP NPOav NPY NVT P4 P450aro P450C17 P450scc

17a,20b-dihydroxyprogesterone 17b-hydroxysteroid dehydrogenase 3b-hydroxysteroid dehydrogenase Corticotropin Androgen receptor Cocaine- and amphetamine-related transcript Chicken gonadotropin-releasing hormone-II Dopamine Dihydrotestosterone 17b-estradiol Estrogen receptor Estrogen receptor-a Follicle-stimulating hormone g-aminobutyric acid Glutamic acid decarboxylase Gonadotropin-releasing hormone-associated peptide Growth hormone Growth hormone-releasing hormone Processing peptide Gonadotropin-inhibiting hormone Gonadotropin-releasing hormone Gonadotropin-releasing hormone receptor Glycoprotein a2 Glycoprotein b5 G-protein-coupled receptor G-protein-coupled receptor 54 (kisspeptin receptor) Glucocorticoid receptor Gonadotropin g-vinyl-GABA Hypothalamicepituitaryegonadal Insulin-like growth factor Immunoreactive Kisspeptin Luteinizing hormone Luteinizing hormone-releasing hormone Mediobasal hypothalamus Median eminence Melatonin Melatonin receptor Mammalian gonadotropin-releasing hormone Million years before the present Nucleus preopticus anteroventralis Neuropeptide Y Nucleus ventral tuberis Progesterone Aromatase 17-hydroxylase 17,20 lyase Cholesterol side-chain cleaving enzyme

PD PI PN POA PPD PR PRL PT RPD sGnRH SS T TH TRH TSH

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Pars distalis Pars intermedia Pars nervosa Preoptic area Proximal pars distalis Progesterone receptor Prolactin Pars tuberalis Rostral pars distalis Salmon gonadotropin-releasing hormone Somatostatin Testosterone Tyrosine hydroxylase Thyrotropin-releasing hormone Thyrotropin

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in neurons and pituitary cells implicated in reproductive functions in rainbow trout: a double immunohistochemical study. Biol. Reprod., 60, 642e650. Thomas, P., Dressing, G., Pang, Y., Berg, H., Tubbs, C., Benninghoff, A., et al. (2006). Progestin, estrogen and androgen G-protein coupled receptors in fish gonads. Steroids, 71, 310e316. Thornton, J., Zehr, J. L., & Loose, M. D. (2009). Effects of prenatal androgens on rhesus monkeys: a model system to explore the organizational hypothesis in primates. Horm. Behav., 55, 633e645. Todo, T., Ikeuchi, T., Kobayashi, T., & Nagahama, Y. (1999). Fish androgen receptor: cDNA cloning, steroid activation of transcription in transfected mammalian cells, and tissue mRNA levels. Biochem. Biophys. Res. Commun., 254, 378e383. Tomy, S., Wu, G. C., Huang, H. R., Dufour, S., & Chang, C. F. (2007). Developmental expression of key steroidogenic enzymes in the brain of protandrous black porgy fish, Acanthopagrus schlegeli. J. Neuroendocrinol., 19, 643e655. Tong, S. K., & Chung, B. C. (2003). Analysis of zebrafish cyp19 promoters. J. Steroid. Biochem. Mol. Biol., 86, 381e386. Trudeau, V. L. (1997). Neuroendocrine regulation of gonadotrophin II release and gonadal growth in the goldfish, Carassius auratus. Rev. Reprod., 2, 55e68. Trudeau, V. L., Sloley, B. D., Wong, A. O., & Peter, R. E. (1993). Interactions of gonadal steroids with brain dopamine and gonadotropin-releasing hormone in the control of gonadotropin-II secretion in the goldfish. Gen. Comp. Endocrinol., 89, 39e50. Trudeau, V. L., Spanswick, D., Fraser, E. J., Lariviere, K., Crump, D., Chiu, S., et al. (2000). The role of amino acid neurotransmitters in the regulation of pituitary gonadotropin release in fish. Biochem. Cell Biol., 78, 241e259. Tsuneki, K., & Gorbman, A. (1975). Ultrastructure of pars nervosa and pars intermedia of the Lamprey, Lampetra tridentata. Cell Tissue Res., 157, 165e184. Tsutsui, K., & Ukena, K. (2006). Hypothalamic LPXRF-amide peptides in vertebrates: identification, localization and hypophysiotropic activity. Peptides, 27, 1121e1129. Tsutsui, K., Bentley, G. E., Ubuka, T., Saigoh, E., Yin, H., Osugi, T., et al. (2007). The general and comparative biology of gonadotropininhibitory hormone (GnIH). Gen. Comp. Endocrinol., 153, 365e370. Twan, W. H., Hwang, J. S., Lee, Y. H., Jeng, S. R., Yueh, W. S., Tung, Y. H., et al. (2006). The presence and ancestral role of gonadotropin-releasing hormone in the reproduction of scleractinian coral, Euphyllia ancora. Endocrinology, 147, 397e406. Uchida, K., Murakami, Y., Kuraku, S., Hirano, S., & Kuratani, S. (2003). Development of the adenohypophysis in the lamprey: evolution of epigenetic patterning programs in organogenesis. J. Exp. Zoolog. B. Mol. Dev. Evol., 300, 32e47. Unniappan, S., & Peter, R. E. (2004). In-vitro and in-vivo effects of ghrelin on luteinizing hormone and growth hormone release in goldfish. Am. J. Physiol. Regul. Integr. Comp. Physiol., 286, R1093eR1101. Unniappan, S., Lin, X., Cervini, L., Rivier, J., Kaiya, H., Kangawa, K., et al. (2002). Goldfish ghrelin: molecular characterization of the complementary deoxyribonucleic acid, partial gene structure and evidence for its stimulatory role in food intake. Endocrinology, 143, 4143e4146.


Vacher, C., Ferriere, F., Marmignon, M. H., Pellegrini, E., & Saligaut, C. (2002). Dopamine D2 receptors and secretion of FSH and LH: role of sexual steroids on the pituitary of the female rainbow trout. Gen. Comp. Endocrinol., 127, 198e206. Vacher, C., Mananos, E. L., Breton, B., Marmignon, M. H., & Saligaut, C. (2000). Modulation of pituitary dopamine D1 or D2 receptors and secretion of follicle stimulating hormone and luteinizing hormone during the annual reproductive cycle of female rainbow trout. J. Neuroendocrinol., 12, 1219e1226. Van Aerle, R., Kille, P., Lange, A., & Tyler, C. R. (2008). Evidence for the existence of a functional Kiss1/Kiss1 receptor pathway in fish. Peptides, 29, 57e64. Vetillard, A., Benanni, S., Saligaut, C., Jego, P., & Bailhache, T. (2002). Localization of tyrosine hydroxylase and its messenger RNA in the brain of rainbow trout by immunocytochemistry and in-situ hybridization. J. Comp. Neurol., 449, 374e389. Vickers, E. D., Laberge, F., Adams, B. A., Hara, T. J., & Sherwood, N. M. (2004). Cloning and localization of three forms of gonadotropinreleasing hormone, including the novel whitefish form, in a salmonid, Coregonus clupeaformis. Biol. Reprod., 70, 1136e1146. Vidal, B., Pasqualini, C., Le Belle, N., Holland, M. C., Sbaihi, M., Vernier, P., et al. (2004). Dopamine inhibits luteinizing hormone synthesis and release in the juvenile European eel: a neuroendocrine lock for the onset of puberty. Biol. Reprod., 71, 1491e1500. Weltzien, F. A., Pasqualini, C., Sebert, M. E., Vidal, B., Le Belle, N., Kah, O., et al. (2006). Androgen-dependent stimulation of brain dopaminergic systems in the female European eel (Anguilla anguilla). Endocrinology, 147, 2964e2973. White, S. A., Kasten, T. L., Bond, C. T., Adelman, J. P., & Fernald, R. D. (1995). Three gonadotropin-releasing hormone genes in one organism suggest novel roles for an ancient peptide. Proc. Natl. Acad. Sci. USA, 92, 8363e8367. Yacobovitz, M., Solomon, G., Gusakovsky, E. E., Levavi-Sivan, B., & Gertler, A. (2008). Purification and characterization of recombinant pufferfish (Takifugu rubripes) leptin. Gen. Comp. Endocrinol., 156, 83e90. Yin, H., Ukena, K., Ubuka, T., & Tsutsui, K. (2005). A novel G proteincoupled receptor for gonadotropin-inhibitory hormone in the Japanese quail (Coturnix japonica): identification, expression and binding activity. J. Endocrinol., 184, 257e266. Yoo, M. S., Kang, H. M., Choi, H. S., Kim, J. W., Troskie, B. E., Millar, R. P., et al. (2000). Molecular cloning, distribution and pharmacological characterization of a novel gonadotropin-releasing hormone ([Trp8] GnRH) in frog brain. Mol. Cell Endocrinol., 164, 197e204. Zhang, D., Popesku, J. T., Martyniuk, C. J., Xiong, H., DuarteGuterman, P., Yao, L., et al. (2009a). Profiling neuroendocrine gene expression changes following fadrozole-induced estrogen decline in the female goldfish. Physiol. Genomics, 38, 351e356. Zhang, D., Xiong, H., Mennigen, J. A., Popesku, J. T., Marlatt, V. L., Martyniuk, C. J., et al. (2009b). Defining global neuroendocrine gene expression patterns associated with reproductive seasonality in fish. PLoS One, 4, e5816. Zohar, Y., Munoz-Cueto, J. A., Elizur, A., & Kah, O. (2010). Neuroendocrinology of reproduction in teleost fish. Gen. Comp. Endocrinol., 165, 438e455.

Chapter 3

Testicular Function and Hormonal Regulation in Fishes Rosemary Knapp and Sharon L. Carlisle University of Oklahoma, Norman, OK, USA

SUMMARY This chapter summarizes our current understanding of teleost testicular and accessory structure function and their hormonal regulation in a phylogenetic context. We review recent work on structure and function, especially with respect to the role of steroid hormones in controlling spermatogenesis. Our review highlights how much of teleost diversity remains effectively unexplored with respect to hormonal regulation of testicular function. We also introduce some examples of within-species variation in male reproductive biology. Future study of such species holds great promise for increasing our understanding of testicular function in general. Throughout, we identify what we consider fruitful avenues for future research, especially those questions where advances in molecular techniques should now allow progress that was not possible in the past. As one example, we note interesting correlations in teleosts between androgen receptor gene evolution and the occurrence of viviparity and alternative male reproductive phenotypes when considered in the light of our current understanding of fish phylogeny.

1. INTRODUCTION There is a great diversity of reproductive modes among teleost fishes (Teleostii) (Breden & Rosen, 1966). This taxon harbors various forms of multiple independent evolutions of sexual and asexual reproduction (Schlupp, 2005), genetic sex determination, and environmental sex determination (see Chapter 1, this volume); both internal and external fertilization (Patzner, 2008); and a variety of mating systems, including some with parental care ranging from solely maternal care to solely paternal care (Blumer, 1979; Ah-King, Kvarnemo, & Tullberg, 2005). With approximately 27 000 species currently recognized (Nelson, 2006; Froese & Pauly, 2009; Eschmeyer & Fong, 2010), it is also the largest vertebrate taxon. Such diversity is both a bonanza and a challenge for researchers. The diversity represents a rich trove from Hormones and Reproduction of Vertebrates, Volume 1dFishes Copyright Ó 2011 Elsevier Inc. All rights reserved.

which to select study species on particular aspects of biology, especially reproductive biology. The diversity represents a challenge because the large number of species means that determining whether or not a species’ particular behavior, morphology, or other characteristic is ‘representative’ for a particular group requires that a substantial number of species be investigated. In addition, sampling only a few species makes inferring how particular traits evolved problematic (see, e.g., Almada & Robalo, 2008). We have several goals for this chapter. First, we summarize our current understanding of testicular and accessory structure function and their hormonal regulation in teleost fishes in an evolutionary context. Readers are referred to Chapters 10e12, this volume, for more detailed discussion of the other groups of fishes. In addition, several excellent reviews of adult testicular function and hormonal regulation in teleosts have appeared recently (Schulz & Miura, 2002; Miura & Miura, 2003; Grier & Uribe Aranza´bal, 2009; Lahnsteiner & Patzner, 2009a; Schulz et al., 2010) that provide more specific details on particular topics than we have space for here. A second goal is to summarize the current state of our understanding in a phylogenetic context, highlighting how much of teleost diversity remains effectively unexplored with respect to hormonal regulation of testicular function. Third, we introduce some examples of intraspecific variation in male reproductive biology, a topic that until recently has not received much attention in the context of testicular function. We believe that more detailed study of intraspecific variation in testicular function and spermatogenesis holds great promise for increasing our understanding of testicular function in general. Throughout, we aim to identify interesting and potentially fruitful avenues for future research, especially those questions where advances in molecular techniques should now make progress possible where it was not in the past. We have chosen to cite reviews at the expense of citing fewer primary papers. These excellent 43

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FIGURE 3.1 Photomicrograph and schematic depiction of spermatogenesis in a seminiferous tubule from zebrafish (Danio rerio, Cyprinidae, Cypriniformes) testes. Progression of spermatogenesis is depicted flowing from the lower left corner around to the upper left corner. Adiff, type A differentiated spermatogonia; Aund), type A undifferentiated spermatogonia (potentially a stem cell); Aund, type A undifferentiated spermatogonia; B (earlyelate), type B spermatogonia; BL, basal lamina; BV, blood vessel; D/MI, diplotene spermatocytes/metaphase I; E1, early spermatids; E2, intermediate spermatids; E3, final spermatids; LE, interstitial Leydig cells; MY, myoid cells; P, pachytene primary spermatocytes; SE, Sertoli cells; S/ MII, secondary spermatocytes/metaphase II; SZ, sperm; Z/L, leptotene/zygotene primary spermatocytes. Reproduced from Schulz et al. (2010) with permission.

reviews on the particular topics provide our readers with a more efficient entry into the literature than would otherwise be possible given our overall goals and space constraints.

2. TESTIS STRUCTURE AND SPERMATOGENESIS: AN OVERVIEW As in all vertebrates, spermatogenesis in fishes begins when spermatogonial stem cells (SSCs) are stimulated by the pituitary gonadotropins (GTHs)dfollicle-stimulating hormone (FSH) and luteinizing hormone (LH) (Kerr, 1995; Schulz & Miura, 2002; Miura & Miura, 2003; Schulz et al., 2010)dto take one of two pathways: self-renewal or maturation. If the SSCs enter the maturation pathway, they undergo a species-specific number of mitotic divisions to produce type A spermatogonia (SG-A), which reside within a cyst. The SG-A then undergo mitotic divisions to form short-lived type B spermatogonia (SG-B). These again undergo mitotic divisions to form primary spermatocytes, which in turn undergo meiosis. The first meiotic division produces secondary spermatocytes, which then undergo the second meiotic division, producing haploid spermatids, which subsequently mature into sperm.

In anamniote vertebrates (fishes and amphibians), spermatogenesis occurs in cysts formed by Sertoli cells. We refer the reader to Leal et al. (2009) and Schultz et al. (2010) for more detailed description of Sertoli cell proliferation and cellular details of spermatogenesis. All germ cells within a cyst originate from a single SSC and develop synchronously, yielding cysts that contain clones of germ cells in the same stage of spermatogenesis. These stages can be distinguished by cell size, nuclear histology and the number of cells per cyst (Schulz et al., 2005; 2010) (Figure 3.1). Depending on the species, spermiation (rupture of the spermatocysts) occurs either after sperm maturation (in externally fertilizing species) or at the spermatid stage, with final maturation occurring in seminal vesicles and often the packaging of sperm into packets called spermatozeugmata (as in many viviparous species with internal fertilization) (Lahnsteiner & Patzner, 2009a). In their recent comprehensive review of spermatogenesis in fishes, Schulz et al. (2010) hypothesize that cystic spermatogenesis exhibited by anamniotes may be more efficient than the noncystic spermatogenesis exhibited by amniote vertebrates. Although the authors do not explicitly state what the selective pressure for this greater efficiency would be, it would seem to correlate with the ubiquitous

Chapter | 3

Testicular Function and Hormonal Regulation in Fishes

(a)

(b)

(c)

FIGURE 3.2 Representative photomicrographs of the major testes types in fishes. (a) Anastomosing tubular testis from the tarpon (Megalops atlanticus, Elopiformes). Anastomosing tubules form continuous, interconnected loops as illustrated by the lobules shaded in light gray. This image from a reproductively active fish shows lobules filled with stored sperm (SP); developing germ cells in spermatocysts are not present, as is characteristic of reproductive males from species that are synchronous spawners. (b) Unrestricted lobular testis from the striped mullet (Mugil cephalus, Mugliformes), collected one month before the spawning season. Spermatocysts containing germ cells in different developmental stages are found in various locations along the length of the lobule. Note in particular that spermatocysts containing spermatogonia (SG) are not limited to the distal end of the lobule. (c) Restricted lobular testis from the Gulf killifish (Fundulus grandis, Cyprinodontiformes). In such testes, spermatogenetic stages progress in developmental sequence along the lobule proximally from the periphery of the testis; spermatocysts containing spermatogonia are restricted to the distal end of lobules. ED, Efferent duct; 1SC, primary spermatocytes; 2SC, secondary spermatocytes; ST, spermatids. Bars ¼ (a) 100 mm, (b, c) 50 mm. Reproduced with permission from Parenti and Grier (2004).

45

use of internal fertilization by the terrestrial amniotes. The use of external fertilization by most anamniote species, which are aquatic or amphibious, is likely to put a higher premium on sperm number than is the case for terrestrial animals. It would be particularly interesting to compare species with cystic spermatogenesis with those in which spermatogenesis is semicystic (e.g., Callichthyidae (Siluriformes), Scorpaenidae (Scorpaeniformes), Blenniidae, and some Gobiidae (both Perciformes)), which typically results in fewer mature sperm at any given time (Lahnsteiner & Patzner, 2009a). Testes contain a germinal epithelium comprised of somatic and germ cells. The arrangement of the germinal epithelium into compartments varies across fish taxa, with three main types identified in a recent comprehensive review of a broad range of species (Parenti & Grier, 2004). Anastomosing tubular testes are found in males from basal lineages such as coelacanths (e.g., the West Indian Ocean coelacanth, Latimeria chalumnae (Coelacanthiformes)), Acipenseriformes, and Cypriniformes and basal euteleosts such as the Salmoniformes (Figures 3.2(a) and 3.3). In these testes, germinal compartments do not terminate at the testis periphery, but rather form highly branched, anastomosing loops. In contrast, lobular testes are defined as those testes that contain some germinal compartments that terminate blindly at the periphery of the testis. Lobular testes are categorized as either unrestricted lobular, where cysts containing spermatogonia are found along the length of the lobules, and restricted lobular, where spermatogonia-containing cysts are found only at the periphery of the lobule (Figure 3.2(bec)). Unrestricted lobular testes are found in neoteleosts with the exception of the Atherinomorpha (Atheriniformes, Beloniformes, and Cyprinodontiformes) and a few scattered species that possess restricted lobular testes (Figure 3.3) (Parenti & Grier, 2004). One particularly interesting exception to the Atherinomorpha-specific distribution of restricted lobular testes is the presence of this testes type in Schindler’s fish (Schindleria praematura) (Schindleriidae, Perciformes) (Thacker & Grier, 2005). The presence of a restricted lobular testis in this species has been hypothesized to be related in some currently unknown way to the fact that the species is paedomorphic (Thacker & Grier, 2005). Despite the large number of fish families surveyed by Parenti and Grier (2004), our understanding of the evolution of testicular development and morphology will undoubtedly be modified in the future as additional species are investigated. Figure 3.3 illustrates the distribution of testicular types in those taxa presented by Parenti and Grier (2004). The topology of this phylogeny is based on Stiassney, Wiley, Johnson, and De Carvalho (2004), with modifications based on Miya, Satoh, and Nishida (2005); Ortı´ and Li (2009); and Broughton (2010). From this figure, one can see just how small the proportion of teleost orders

46


Teleostei

Otocephala

Neopterygii

#SP

1

12

Acipenseriformes

2

28

Lepisosteiformes

1

7

Amiiformes Osteoglossomorpha

1 7

1 213

Elopomorpha

26

972

Clupeiformes

6

392

Gonorynchiformes

4

37

6 25

3,886 2,389

Cypriniformes Characiformes AR b Losses

Clupeocephala

AR Duplication

#FAM Polypteriformes

Gymnotiformes

NOTES

5

1

11 2

A2

5

158

38

3,357

5

A1

Salmoniformes Esociformes

1 2

202 12

8

A1 A1

Osmeriformes

12

310

6

202

4 16

421 258

Myctophiformes

2

256

Lampriformes

7

26

Polymixiiformes

1

10

Ateleopodiformes Lophiiformes

1 18

13 336

Gadiformes

Siluriformes

Argentiniformes Euteleostei

Stomiiformes Aulopiformes

Acanthopterygii

10

607

Percopsiformes

3

9

Ophidiiformes Zeiformes

5 6

476 33

Atheriniformes

11

394

5

258

1

V1 A3

10

1,170

1

V3 A4

Mugiliformes Beryciformes

1 7

73 158

Stephanoberyciformes

5

94

Synbranchiformes

3

113

1

A1

Gobiesociformes Batrachoidiformes

1 1

158 80

Scorpaeniformes

35

1,572

5

29

5 10

361 431

11

784

5

162 10,598

23

Beloniformes Cyprinodontiformes

AR b Neofunctionalization

ESP

Percomorpha

Gasterosteiformes Syngnathiformes Tetraodontiformes Pleuronectiformes Perciformes

1 V3 2

A1 V2 1

A2 A3 V4 A15

FIGURE 3.3 Distribution of major testes types, species studied for hormonal regulation of testicular function, and families with viviparity or male alternative reproductive tactics across actinopterygian orders or superorders. Topology is based on Stiassney, Wiley, Johnson, and De Carvalho (2004), with modifications based on Miya, Satoh, and Nishida (2005); Ortı´ and Li (2009); and Broughton (2010, and personal communication), although the phylogeny of the Acanthopterygii is in a state of flux; line lengths do not reflect degree of divergence. Black bars indicate whole genome duplication events. The three major testes types as reported in Parenti and Grier (2004) are indicated by font: anastomosing tubular (underlined), restricted lobular (bold), unrestricted lobular (italicized bold). Numbers of families (#FAM) and species in each family (#SP) are from Eschmeyer & Fong (2010). Number of species in which endocrine aspects of testicular function have been studied (ESP) is based on a search of Web of Science in August 2009 using the terms “fish, teleost, testes, testis, hormone(s)” in various combinations. Also indicated are those orders where at least one species has been documented to exhibit viviparity (V) (from Wourms, 1981; Contreras-Balderas, 2005) or alternative reproductive tactics (A) (from Knapp & Neff, 2008), with the number of families represented indicated. Timing of events in androgen receptor (AR) gene evolution is from Douard et al. (2008).

Chapter | 3


covered by their extensive survey was. Indeed, since publication of their review, an apparently previously undescribed testicular type has been described in Atlantic cod (Gadus morhua, Gadidae, Gadiformes) (Almeida, Kristoffersen, Taranger, & Schulz, 2008). In this species, the mature testis contains many lobes, with each lobe containing a gradient of progressively mature germ cells. The description is reminiscent of restricted lobular testes, but additional work is required to determine the extent of the similarity. Another example of an apparently different type of testis has been documented in two species of pipefish (Syngnathidae, Syngnathiformes). In these species, where males fertilize eggs that females transfer to them for brooding, the testis is a single hollow tube that does not form the cysts typically seen in other fishes (Carcupino et al., 1999; Watanabe, Hara, & Watanabe, 2000). Spermatogenesis in these species also appears to be unusual, as is ovarian structure in females (Begovac & Wallace, 1987). Additional work on these two species, and other syngnathids, is certain to provide important insight into testicular development and spermatogenesis in this group with highly unusual reproductive behavior. In addition to variation in testicular morphology across fish species, the morphology and other characteristics of the sperm themselves also vary considerably. In fact, the variation is such that sperm ultrastructural features have been used as characters in phylogenetic analyses of fish evolution and systematics (Jamieson 1991; 2009; Lahnsteiner & Patzner, 2009b). Several sculpin species (Cottidae, Scorpaeniformes) are known to produce both typical flagellated haploid sperm (eusperm) and atypical sperm (parasperm) that are incapable of fertilizing eggs (Hann, 1930; Quinitio, Takahashi, & Goto, 1988). In one sculpin species, these atypical sperm may help increase the distance that the milt can travel to increase fertilization success and may benefit males in sperm competition (Hayakawa, Akiyama, Munehara, & Komaru, 2002). Another example of intraspecific sperm dimorphism comes from some syngnathids (Syngnathidae, Syngnathiformes). In both the seaweed pipefish (Syngnathus schlegeli) (Watanabe, Hara, & Watanabe, 2000) and yellow seahorse (Hippocampus kuda) (Van Look, Dzyuba, Cliffe, Koldewey, & Holt, 2007), sperm fall into one of two morphologically distinct groups. However, the contribution, if any, of hormones or other chemical messengers to such species-level variation is currently unknown. Additional inter-male variation in sperm characteristics has been described in some species, but again our understanding of endocrine contributions to this variation is still very poor (see Section 7).

3. TESTICULAR HORMONES As in other vertebrates, fish testes are organs of both gamete and hormone production. Here we briefly describe

47

the major endocrine molecules known to be produced by testes in fishes.

3.1. Steroids Steroid synthesis by the testes of male teleosts proceeds, as in other vertebrates, from the conversion in the Leydig cells of cholesterol through a series of enzymatic steps to pregnenolone and then via different sub-pathways to various androgens, estrogens, progestogens, and corticosteroids (Norris, 2007). The particular steroids produced are determined by the presence or absence of particular enzymes that act at specific sites on the steroid ring structure (reviewed in Young, Kusakabe, Nakamura, Lokman, & Goetz, 2005). The number of potential steroids is quite large. Whether a particular steroid is detectable depends both on its absolute level in the tissue being assayed and on the tools used to conduct the assay. As has been highlighted by other authors (Kime, 1993; Scott, Sumpter, & Stacey, 2010), our knowledge of teleost reproductive hormones may be biased by reliance on commercial antisera that have often unknown cross-reactivity with the various fish steroids, and use of these antibodies must be validated for each species. Kime (1993) also cautioned that the availability of particular antisera determines what one can find in the plasma or various tissues of nonmammalian speciesd you can only find something if your method can detect it. This caution is as important today as it was 15 years ago, given the increasing availability of commercial enzymelinked immunosorbent assay (ELISA) and radioimmunoassay (RIA) kits that, with a few notable exceptions, have been developed for laboratory mammals and humans. Methods that do not depend on antibodies, such as liquid chromatography-mass spectrometry (LC-MS) and high pressure liquid chromatography (HPLC), have been helpful in identifying additional steroids of potential interest, but these methods also require that suitable standards are available. With the above caveats in mind, the major circulating androgens in male teleosts are testosterone (T), 11-ketotestosterone (11-KT), 11b-hydroxytestosterone (OHT), and 11b-hydroxyandrostenedione (OHA) (Kime, 1993; Borg, 1994). Teleosts produce a number of additional steroids, which are found in the plasma of various species (Kime, 1993). Teleost testes also produce steroid conjugates, particularly glucuronides, which are more commonly produced in the liver in other vertebrates (Fostier, Jalabert, Billard, Breton, & Zohar, 1983; Kime, 1993). Most antisera have low or unknown cross-reactivity with these steroid metabolites. Fostier et al. (1983) reviewed the biochemical studies then available on the specific steroidogenic pathways for androgen synthesis. Although methods differed across species, there was evidence of species-level variation in the

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particular pathway(s) taken to produce 11-KT. These authors summarized several different pathways that have been hypothesized for testicular androgen synthesis. The favoring of particular pathways could be related to variation in substrate concentrations, which may vary at different points during spermatogenesis and/or reproductive season (e.g., Abdullah & Kime, 1994). Genomic and molecular methods should help identify the particular steroidogenic pathways used in particular species at particular points of spermatogenesis. For example, Kusakabe, Nakamura, Evans, Swanson, and Young (2006) found that expression of steroidogenic acute regulatory (StAR) protein and 17a-hydroxylase/17,20b-lyase (P450c17) are likely to be the major influences on overall steroidogenic output in rainbow trout. Such knowledge is not just important to molecular endocrinologists. Knowledge of the particular enzymes involved in steroid synthesis and metabolism is critical for a complete understanding of the evolution of steroid signaling pathways, evolution of mechanisms associated with variation in sperm characteristics (see Section 7), coordination of spermatogenesis with reproductive behavior such as courtship and paternal care, and understanding various tradeoffs, e.g., between androgen-dependent spermatogenesis and androgensensitive immune function (e.g., Ros, Ferreira, Santos, & Oliveira, 2006). Other steroids produced by the testes and/or present in milt have received less attention than have the androgens, although that is changing. Estradiol (E2), some of testicular origin, is increasingly being recognized as critical for certain stages of spermatogenesis (see below). The progestogens 17,20b-dihydroxypregn-4-en-3-one (17,20bDHP) and 17,20adihydroxypregn-4-en-3-one (17,20aDHP) have been found in testicular incubations or plasma of several species (reviewed in Kime, 1993; Scott, Sumpter, & Stacey, 2010). 5a- and 5b-reduction of progestogens (and androgens) has also been documented in vitro. Corticosteroids also have been detected in the testes of goldfish (Carassius auratus, Cyprinidae, Cypriniformes), produced via enzymatic conversion of 17-hydroxyprogesterone (Kime & Scott, 1993). The precursors for steroids produced by the testes need not all come from the testes themselves (Fostier, Jalabert, Billard, Breton, & Zohar, 1983); plasma-borne interrenal and even liver products may be important substrates for testicular androgen production (see Kime, 1978).

3.2. Additional Hormones and Signaling Molecules More recently, additional local testicular signaling molecules have received attention. One of these is the transforming growth factor-b (TGFb) superfamily. Various members of this family have been found to be expressed


in fish testicular tissue. For example, activin Bb is expressed in Sertoli cells of the Japanese eel (Anguilla japonica, Anguillidae, Elopomorpha) and is upregulated by treatment with human chorionic gonadotropin (reviewed in Nagahama, 1994). Activin Bb also stimulates goldfish FSHb but suppresses LHb expression in the pituitary (reviewed in Ge, 2005). Insulin-like growth factors-I and-II (IGF-I and IGF-II) and their receptors also are present in the testes (e.g., Le Gac, Loir, LeBail, & Ollitrault, 1996; Perrot & Funkenstein, 1999). Testicular growth hormone (GH) receptors are also known (Gomez, Loir, & Le Gac, 1998; Ma et al., 2007). However, our understanding of the role of these molecules in steroidogenesis and spermatogenesis is still in its infancy. This will fortunately change with the increased availability of various molecular techniques and genomic approaches.

3.3. Genomic Approaches Genomic approaches and associated molecular and transgenic techniques are increasing the identification of molecules known to be important in spermatogenesis and steroidogenesis. Better understanding of the latter process is critical for understanding how sperm production is coordinated with reproductive behavior. To date, gene expression studies of testicular development and function in fishes have of necessity focused on a limited number of species: rainbow trout (Oncorhynchus mykiss, Salmonidae, Salmoniformes) (Mazurias, Montfort, Delalande, & Le Gac, 2005; Rolland et al., 2009), Atlantic salmon (Salmo salar, Salmonidae, Salmoniformes) (Maugars & Schmitz, 2008), Japanese eel (Anguillidae, Elopomorpha) (Miura & Miura, 2001; Miura, Kuwahara, & Miura, 2007), goldfish (Cyprinidae, Cypriniformes) (Marlatt et al., 2008), zebrafish (D. rerio, Cyprinidae, Cypriniformes) (Zeng & Gong, 2002; Li et al., 2004; Santos et al., 2007; Sreenivasan et al., 2008; Small, Carney, Mo, Vannucci, & Jones, 2009), fathead minnow (Pimephales promelas, Cyprinidae, Cypriniformes) (Kane et al., 2008), medaka (Oryzias latipes, Adrianichthyidae, Beloniformes) (Lo, Zhang, Hong, Peng, & Hong, 2008), bluefin tuna (Thunnus thynnus, Scombridae, Perciformes) (Chini et al., 2008), and sea bass (Dicentrarchus labrax, Moronidae, Perciformes) (Vinãs & Piferrer, 2008). A number of other fish species are being studied with microarrays but testicular function and spermatogenesis have not been the focus of these studies (e.g., Miller & Maclean, 2008; Renn, Aubin-Horth, & Hofmann, 2008). In addition, germ cells marked with vasa-driven fluorescent proteins have been developed for a few species (e.g., rainbow trout (Yoshizaki, Takeuchi, Sakatani, & Takeuchi, 2000; Yano, Suzuki, & Yoshizaki, 2008); medaka (Tanaka, Kinoshita,

Chapter | 3


Kobayashi, & Nagahama, 2001); zebrafish (Krøvel & Olsen, 2004)). Such studies have increased, and will continue to increase, our understanding of signaling molecules involved in spermatogenesis. Of these species, however, none exhibits paternal care, thus in one sense representing ‘simple’ cases of spermatogenesis where there has not been selection for varying, competing functions of androgens (for example). How comparable will results be with those from species such as the threespined stickleback (Gasterosteus aculeatus, Gasterosteidae, Gasterosteiformes) where males do build nests and provide paternal care? The stickleback genome is currently being sequenced and promises to help expand our understanding of such questions in the near future. Another example of genomic approaches relevant to our topic is the use of DNA sequencing to understand the evolution of particular genes of interest. For example, two recent studies have explored the evolution of androgen receptor (AR) genes in fishes and jawed vertebrates (Douard et al., 2008; Ogino, Katoh, Kuraku, & Yamada, 2009). These authors have documented an ancient duplication of the ancestral AR gene. Douard et al. (2008) also identified two subsequent, independent losses of the ARb gene and proposed that neofunctionalization of the ARb gene (i.e., acquisition of new function(s) relative to the ARa gene) may have played a role in the great diversification of the percomorphs. It will be very interesting to see whether or how specific base-pair substitutions in the ligand- and DNA-binding domains of the ARb gene within the percomorphs (Douard et al., 2008) relate to variation in steroidogenesis, testis morphology, and/or spermatogenesis. When we placed the timing of these events in ARb gene evolution on our current understanding of teleost phylogeny (Figure 3.3), we were struck by intriguing correlations with the distribution of viviparity and the expression of alternative male reproductive tactics. The distribution of viviparity across actinopterygians (Wourms, 1981; Contreras-Balderas, 2005) appears after the neofunctionalization of ARb proposed by Douard et al. (2008). Exploration of whether specific substitutions in the ARb gene are correlated with the several independent evolutions of viviparity promises to be a very rich area for future study. For example, many aspects of reproductive biology and behavior are modified in these species and development of the intromittent organ, in particular, is known to be under the control of androgens in several species (see Uribe Aranza´bal, Grier, De la Rosa Cruz, & Garcıá Alarcoń, 2009). Similarly, the distribution of families in which alternative male reproductive tactics have been documented (Figure 3.3) (Knapp & Neff, 2008) shows an intriguing association with important events in ARb gene evolution proposed by Douard et al. (2008) and suggests another fruitful area for future research.

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4. ENDOCRINE REGULATION OF TESTIS STRUCTURE AND FUNCTION The endocrine basis of spermatogenesis in fishes has received much attention since the beginning of the 21st century, although mainly in commercially important species such as salmonids, catfishes, and eels (reviewed in Nagahama, 1994; Schulz & Miura, 2002; Miura & Miura, 2003; Young, Kusakabe, Nakamura, Lokman, & Goetz, 2005; Schulz et al., 2010). We briefly summarize some major features. The two GTHs have generally differing roles in spermatogenesis: FSH during early stages of spermatogenesis and LH during the final stages of maturation. Progestins, 11-KT, and E2 are all key sex steroids, but at different stages of spermatogenesis. Low doses of E2 stimulate the spermatogonial self-renewal pathway, 11-KT controls initiation of the maturation pathway, and 17,20bDHP is critical for initiating meiosis and also for spermiation (Figure 3.4) (Miura, Yamauchi, Takahashi, & Nagahama, 1992; T. Miura, Higuchi, Ozaki, Ohta, & C. Miura, 2006; Schulz & Miura, 2002; T. Miura & C. Miura, 2003; Scott et al., 2010; Schulz et al., 2010). Testosterone, in contrast, can interfere with spermatogenesis (reviewed in Schulz & Miura, 2002). The relationship of both plasma and testicular androgen concentrations to spermatogenesis is complex, and many factors such as sensitivity to androgens and metabolic conversion of androgens have the potential to affect spermatogenesis (Young et al., 2005). The existence of two different ARs in some species of teleosts, first identified in the Atlantic croaker (Micropogonias undulatus, Sciaenidae, Perciformes (Sperry & Thomas, 1999)) and rainbow trout (Takeo & Yamashita, 1999), likely explains the differences in physiological activity of T and 11-KT, with 11-KT’s effects probably mediated through one AR type and T’s effects mediated via both AR types. Perhaps steroidogenic enzymes also play a role in determining which AR is activated by influencing local levels of these two ligands. Additional chemical messengers can influence testicular functions in fishes. Insulin-like growth factor-I and IGF-II act as local mediators of the actions of GH and GTHs and also act directly on the germ cells to stimulate proliferation (Loir & Le Gac, 1994; Le Gac et al., 1996; Perrot & Funkenstein, 1999). The possible roles of thyroid hormones in reproduction are discussed in Chapter 5, this volume. Stress hormones, such as cortisol, can alter male reproductive function (see Milla, Wanga, Mandikia, & Kestemonta, 2009) (see also Chapter 6, this volume). For example, Ozaki et al. (2006) found that lower doses of cortisol had positive effects on spermatogonial proliferation, whereas higher doses were inhibitory. Stimulatory effects are likely related to cortisol’s ability to trigger mobilization of energy reserves. Cortisol’s inhibitory effects over some range of levels may be ameliorated

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Leydig Cell

FSH T

11-KT

E2

Sertoli Cell

Spermatogonial Renewal Factor

LH

P450c17 More 17a-OHP4 available

AMH

17,20bP

Activin B 17,20bP

1n

1n

1n

1n

1n

1n

1n

1n

1n

1n

1n

1n

2n 2n 2n

2n

2n

2n 2n

renewal

proliferation

meiosis

spermiogenesis

FIGURE 3.4 Schematic illustration of major hormonal influences on spermatogenesis in teleosts. The stages of spermatogenesis are delineated at the bottom of the figure and correspond to the various germ cell stages. The ploidy level of the germ cell stages are indicated by 2n or 1n. Circles with arrows indicate points of cell proliferation (spermatogonial stem cells and type B spermatogonia). The dotted line through the Sertoli cell separates control of renewal from control of proliferation. 11-ketotestosterone is represented outside the Leydig cell to reflect additional non-testicular sites of synthesis. 17aOHP4, 17a-hydroxyprogesterone; 17,20bP, 17,20b-dihydroxypregn-4-en-3-one; AMH, anti-Mu¨llerian hormone; E2, estradiol; FSH, follicle-stimulating hormone; 11-KT, 11-ketotestosterone; LH, luteinizing hormone; P450c17, 17a-hydroxylase; T, testosterone.

via the enzyme 11b-hydroxysteroid dehydrogenase (11b-HSD), which can inactivate glucocorticoids at several tissues, including the testes (reviewed briefly in Knapp, 2003; Perry & Grober, 2003). Kisspeptins (Kps), neuropeptides that have been implicated as a sort of master switch for turning on reproduction in mammals (Kauffman, Clifton, & Steiner, 2007), are of particular interest in fishes (for a discussion of Kps in fishes, see Chapter 2, this volume). Finally, in addition to naturally occurring compounds, an increasing number of human-produced compounds are being documented to interfere with male reproductive function. For a detailed discussion of these endocrine-disrupting compounds and their effects we refer the reader to Chapter 13, this volume. Many studies of endocrine control of spermatogenesis are conducted in vitro, with testicular slices incubated with different molecules. Another common approach is to correlate circulating levels of various hormones with testicular histology, especially for seasonal breeders such as salmonids. However, plasma levels of hormones do not necessarily reflect the levels of those hormones in the testes themselves. For example, Cavaco, Vischer, Lambert, Goos,

and Schulz (1997) documented that there was a mismatch between circulating and testicular levels of OHA and 11-KT in the African or sharptooth catfish (Clarias gariepinus, Clariidae, Siluriformes). They found that both liver and seminal vesicles were important sites of 11-KT synthesis, and that extra-testicular conversion of OHA accounts for the mismatch observed between plasma and testes. Such findings further complicate our understanding of an inherently complicated process. A further complicating factor is that the testiseblood barrier in fishes apparently varies across the spermatogenetic cycle, unlike the case in mammals (reviewed in Bergmann, Sehindelmeiser, & Greven, 1984; Batlouni, No´brega, & França, 2009). Only cysts containing haploid germ cells have tight junctions joining the Sertoli cells, thus protecting these spermatogenetic stages from chemicals circulating in the plasma. In contrast, meiotic germ cells (SG-A and SG-B) in fishes are more exposed to potential perturbation by changes in chemical messengers and/or exposure to endocrine disruptors. Additional research into the seminiferous epithelium structure should provide important information relative to our understanding of the

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control of germ cell survival and final morphology and performance.

5. TEMPORAL ASPECTS OF TESTICULAR FUNCTION 5.1. Development from Undifferentiated Gonad to Functional Testis The development of testes from the undifferentiated gonad in teleosts is discussed in detail in Chapter 1, this volume, and gonadal differentiation in sex-changing fishes is discussed in Chapter 8, this volume. Here, we briefly discuss one aspect of testicular development associated with the onset of spermatogenesis, both initially, and yearly for annually breeding species: the number of mitotic divisions by SSCs. This feature ultimately determines the number of sperm produced within a spermatocyst. There are two ways of increasing the number of sperm: increasing the number of spermatogonia and increasing the number of rounds of mitosis that spermatogonia undergo before initiation of meiosis (Schulz et al., 2010). The number of mitotic divisions undergone by SSCs varies across species in vertebrates, including fishes. An understanding of control of this step in spermatogenesis is needed to fully understand how sperm numbers are regulated, both at sexual maturation and seasonally for species that undergo seasonal regression and recrudescence. Ando, T. Miura, Nader, C. Miura, and Yamauchi (2000) developed and validated a method for estimating the number of mitotic divisions in fish testes based on counting the number of spermatocytes or spermatids within cysts. More recently, Leal et al. (2009) used a more labor-intensive stereological method to obtain a very accurate determination of the number of spermatogonial divisions. Both methods make use of the cystic nature of fish spermatogenesis. The arrangement of fish germ cells in clonal cysts also allows the use of laser-capture microdissection methods to determine stage-specific gene expression (Vinãs & Piferrer, 2008; Rolland et al., 2009). Such genomic approaches promise to greatly increase the number of chemical messengers implicated in controlling the various aspects of spermatogenesis, and hold particular promise for understanding transitions between various developmental stages during spermatogenesis. In addition, recent results indicating that centriole behavior during meiosis determines the ability of stem cells to regenerate (Cheng et al., 2008; Wang et al., 2009) suggest a potentially promising line of future research in terms of understanding species differences in the number of spermatogonial renewal generations (and hence number of sperm per cyst). However, the role, if any, of centriole behavior in interspecific variation in fish spermatogenesis is currently unknown.

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5.2. Release of Sperm The timing of sperm release is under neuroendocrine control, with pheromones being major stimuli (Kobayashi, Sorensen, & Stacey, 2002) (see also Chapters 2, 7, and 9, this volume). Neuroanatomical and neuroendocrine control of sperm (and egg) release has been studied best in goldfish. Electrical stimulation studies have identified brain areas that trigger sperm (and egg) release in several species (Demski, Baurer, & Gerald, 1975; Demski, 1983). The neuroanatomical pathways controlling sperm release are similar across several percomorph and cypriniform species, involving contractions of the sperm ducts, but apparently differ from the pathways that control sperm release in salmonids (summarized in Dulka & Demski, 1986). Testicular contractions do not seem to be required for sperm release; their function may be to transport sperm from testes to sperm ducts, effectively ‘loading’ them for upcoming spawning bouts (Dulka & Demski, 1986). Such a loading mechanism varies across males within a single species depending on prior experience or exposure to females or other males (e.g., Stacey, Fraser, Sorenson, & Van Der Kraak, 2001; Bozynski & Liley, 2003; Aspbury & Gabor, 2004). Courtship and spawning induce elevations in GTH and androgen levels in male goldfish (Kyle, Stacey, Peter, & Billard, 1984; Sorensen, Pinillos, & Scott, 2005). Two pheromones, prostaglandin F2a (PGF2a) and 17,20b-DHP are important in stimulating milt production, but do so via separate mechanisms (Zheng & Stacey, 1996; 1997) (see also Chapter 7, this volume). The effects of 17,20b-DHP appear to require release of pituitary LH, whereas the effect of the spawning stimulus from a female is LH-independent. The ability of 17,20b-DHP to increase LH and milt volume could be blocked with dopamine (DA) D-2 receptor agonists but spawning-induced responses could not (Zheng & Stacey, 1997). More recently, PGF2a increased salmon gonadotropin-releasing hormone (sGnRH) mRNA levels in the telencephalon and cerebellum of male goldfish, whereas 17a,20b-DHP increased plasma androstenedione (AND) levels but had no effect on sGnRH mRNA levels (ChungDavidson, Rees, Bryan, & Li, 2008). Further exploration of individual variation among males in their sensitivity to these pheromones and other stimuli from the female could help provide insights into the physiological mechanisms related to sperm competition, which constitute a major contributor to a male’s reproductive success in many species (see discussion of within-species variation in Section 7).

5.3. Seasonal Aspects of Testicular Function Fish species vary as to when during the year active spermatogenesis occurs (Munro, Scott, & Lam, 1990; LeGac & Loir, 1999). For some, active spermatogenesis occurs during

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the summer; for others it occurs in the spring. Yet other species begin spermatogenesis in the fall and complete it in the spring. Many marine species reproduce on lunar, semilunar or tidal schedules (Robertson, 1991; Takemura, Rahman, Nakamura, Park, & Takano, 2004). Seasonal changes in photoperiod are also accompanied by changes in water temperature, thus making it a challenge to determine the separate contributions of these two factors (Pankhurst & Porter, 2003). Does hormonal control of reproduction and testis function vary with the season of active spermatogenesis or with the time of actual spawning? Our understanding is still very poor in these areas, due in no small part to the difficulty of conducting the required studies. Many studies of photoperiod effects on fish reproduction focus on ovulation rather than spermatogenesis (see reviews by Pankhurst & Porter, 2003; Takemura et al., 2004). However, a series of studies on rabbitfishes (Siganidae, Perciformes) has documented changes in steroidogenesis and spermatogenesis in association with lunar cycles, with species-level variation in the lunar phase with which spermiation is synchronized (reviewed in Takemura et al., 2004). Studies on effects of photoperiod have focused on species that are important for aquaculture, especially various salmonids and sea bass (e.g., Bromage, Porter, & Randall, 2001). The methods used in these studies, such as exposure to continuous light, may not reflect the actual role of photoperiod in free-living fishes. None-the-less, such studies do provide helpful information on potential physiological mechanisms by which photoperiod might influence testicular function. In general, longterm exposure to continuous light inhibits initiation of reproduction, likely through inhibition of melatonin (MEL) secretion by the pineal (e.g., Oliveira, Ortega, Lo´pezOlmeda, Vera, & Sańchez-Va´zquez, 2007). Amano et al. (2000) showed that exogenous MEL mimicked the stimulatory effects of short photoperiod on precocious testicular development in masu salmon (Oncorhynchus masou, Salmonidae, Salmoniformes) and also stimulated the production and release of pituitary FSH but not LH. Both exogenous MEL and constant darkness also induced precocious maturation of catla (Gibelion (Catla) catla, Cyprinidae, Cypriniformes) testicular fragments, but only during the preparatory period of the annual cycle (Bhattacharya, Chattoraj, & Maitra, 2007). Many questions remain regarding the mechanisms by which seasonal and lunar cycle light cues are transduced to influence testicular function. One might hypothesize that MEL would be more important in regulating spermatogenesis in species living at latitudes where changes in photoperiod are more pronounced. Similarly, it will be interesting to determine whether MEL plays a greater role in species that typically spawn at dawn or dusk compared to species that spawn throughout the day. Finally, what is the role of other chemical messengers


such as Kps and steroid hormones in transducing various environmental signals to influence spermatogenesis (e.g., Bornestaf, Antonopoulou, Mayer, & Borg, 1997; Migaud, Davie, & Taylor, 2010)?

6. ACCESSORY GONADAL STRUCTURES Male fish exhibit tremendous variety in their efferent duct systems, from testes that essentially empty into the cloaca, to partially shared spermatic and renal ducts, to completely separate spermatic and renal duct systems (for details see Lahnsteiner & Patzner, 2009a). In teleosts, the renal and genital systems are completely separate (Nagahama, 1983; Lahnsteiner & Patzner, 2009a). In most species, the testes are located cranially from the genital papilla, with testicular main ducts continuing from the testes into paired spermatic ducts at the caudal end of the testes (Lahnsteiner, Nussbaumer, & Patzner, 1993; Lahnsteiner, Patzner, & Weismann, 1993; 1994; Lahnsteiner, 2003). These paired ducts join to form a single sperm duct that empties into the genital papilla in fish with external fertilization, or, in viviparous fish, into storage structures such as the epididymal gland until copulation (Fishelson, Gon, Holdengreber, & Delarea, 2006). Additional structures such as the genital papilla (of external fertilizers) and modified anal fins (the gonopodia of internal fertilizers) are also important for successful male reproduction in many fishes via their role in sperm transfer (Evans & Meisner, 2009). These secondary sexual characteristics are known in several species to be under the control of androgens (e.g., Turner, 1960; Carlisle et al., 2000; reviewed in Norris, 2007). There are three types of accessory gonadal structure: testicular glands, testicular blind pouches, and seminal vesicles (Lahnsteiner & Patzner, 2009a). Testicular glands are located on the ventral side of the testis and can be thought of as modified testicular efferent ducts. Testicular blind pouches are tube-like evaginations of the spermatic duct. Seminal vesicles are lobular glands that empty into the spermatic ducts, without sphincter muscles. Various teleost families are known to have one or more types of accessory gonadal structure (summarized in Seiwald & Patzner, 1989; Miller, 1992; Rasotto, 1995; Mazzoldi, Lorenzi, & Rasotto, 2007; Lahnsteiner & Patzner, 2009a). Some Blenniidae (Perciformes) have testicular glands and testicular blind pouches. In contrast, testicular glands but no blind pouches are found in the Dactyloscopidae, Tripterygiidae, and Labrisomidae, and in Clinidae that exhibit external fertilization (all Perciformes). Chaenopsidae (Perciformes) have testicular glands and seminal vesicles, and some Gobiidae (Perciformes) have seminal vesicles and steroid-producing glands attached to the testes, which are not homologous to the testicular glands of Blenniidae. Seminal vesicles are found in several siluriform families (Auchenipteridae,

Chapter | 3


Bagridae, Callichthyidae, Clariidae, Heteropneustidae, Ictaluridae, Pimelodidae, Siluridae), Batrachoididae (Batrachoidiformes), and some perciform families (Chaenopsidae, Gobiidae, Eleotridae, and some Blenniidae).

6.1. Testicular Glands and Testicular Blind Pouches The size of testicular glands varies among species (Lahnsteiner, Richtarski, & Patzner, 1990; Richtarski & Patzner, 2000). In species with testicular glands, such as Blenniidae, spermatids do not fully mature into sperm within the testis, but rather complete maturation in seminiferous lobules within the testicular glands (Lahnsteiner & Patzner, 2009a). The glands contain steroidogenic cells homologous to Leydig cells and they also provide nutrition (in the form of glycogen) to developing spermatids. They regulate spermatid differentiation, phagocytose excess spermatids at end of the spawning period, and store lipids (Lahnsteiner & Patzner, 2009a). The glands also secrete mucopolysaccharides and sialomucins into the seminal fluid (reviewed in Lahnsteiner & Patzner, 2009a). These substances increase the viscosity of seminal fluid, which aids in the agglutination of sperm. They may also play a role in fertilization in some species by helping to adhere a layer of seminal fluid, from which sperm are released, onto the substrate on which females deposit eggs (e.g., Ota, Marchesan, & Ferrero 1996; Scaggiante, Mazzoldi, Petersen, & Rasotto, 1999). Unlike testicular glands, testicular blind pouches do not contain sperm (Lahnsteiner, Nussbaumer, & Patzner, 1993). They do, however, contain 3b-steroid dehydrogenase, glucose-6-phosphate dehydrogenase, and uridinediphosphoglucose dehydrogenase, indicating that they may participate in steroid glucuronide synthesis, possibly for use as pheromones.

6.2. Seminal Vesicles Seminal vesicles are glandular outgrowths of the sperm duct. They vary among species with respect to size, morphology, and number of lobes (Van Tienhoven, 1983; Lahnsteiner & Patzner, 2009a), and within species among discrete male phenotypes that exhibit alternative mating tactics (Scaggiante, Grober, Lorenzi, & Rasotto, 2004). Near their junction with the testes, they also may contain spermatocysts in various developmental stages (Fishelson, Vanvuren, & Tyran, 1994; Singh & Joy, 1998; Lahnsteiner & Patzner, 2009a).

6.2.1. Seasonal variation Seminal vesicles exhibit morphological variation throughout the reproductive season (Chowdhury & Joy,

53

2007). They are relatively small during resting and early preparatory phases, increase in size during the mid-preparatory phase, and peak in size during prespawning and spawning phases. They regress again during the postspawning phase. Secretory activity has been studied in catfish (the stinging catfish, Heteropneustes fossilis, Heteropneustidae; the walking catfish, Clarias batrachus, Clariidae; both Siluriformes) throughout this seasonal cycle. Seminal vesicle somatic index and seminal vesicle secretory activity both correlate positively with gonadosomatic index (GSI) (the mass of the gonads divided by the mass of the total body) and testicular secretory activity (Senthilkumaran & Joy, 1993; Singh & Joy, 1999). In addition, steroidogenic activity of seminal vesicles and testes are positively correlated, suggesting that androgens promote secretory activity and that their decline corresponds with the quiescence of the seminal vesicles (Chowdhury & Joy, 2007).

6.2.2. Steroidogenesis The interstitial cells of the seminal vesicles together with the testes originate from the genital ridge (Schoonen & Lambert, 1986; Fishelson et al., 1994) and are considered to be homologous to the testicular Leydig cells (Chowdhury & Joy, 2007). These steroidogenic interstitial cells contain steroid precursors as well as enzymes of steroid hydroxylation (Fishelson et al., 1994; Singh & Joy, 1998). High activity of 3b-steroid dehydrogenase, glucose-6-phosphate dehydrogenase, and uridine-diphosphoglucose dehydrogenase, and reduced activity of NADH dehydrogenase, suggest that steroids and their glucuronides are being synthesized (Nayyar & Sundararaj, 1969; Schoonen, Lambert, & Vanoordt, 1988; Van den Hurk, Schoonen, Van Zoelen, & Lambert, 1987; Asahina, Suzuki, Hibiya, & Tamaoki, 1989; Lahnsteiner, Richtarski, & Patzner, 1990; Singh & Joy, 1998). In the round goby (Neogobius melanostomus, Gobiidae, Perciformes), seminal vesicles synthesize various other androgens from AND in vitro (Jasra et al., 2007).

6.2.3. Hormonal regulation Seminal vesicles rely on androgens for their maintenance and secretory activity (Chowdhury & Joy, 2007). Studies in several catfishes indicate that these androgens are produced by the interstitial cells and act locally to promote secretory activity by the epithelial cells (Singh & Joy, 1998; Viveiros, Eding, & Komen, 2001). The mechanisms of androgen regulation vary among species. In catfishes, castration leads to compensatory hypertrophy and hyperactivity of the seminal vesicles, an effect that can be suppressed by treatment with anti-androgens (e.g., Sundararaj & Nayyar, 1969a; 1969b; Senthilkumaran & Joy, 1993; reviewed in Chowdhury & Joy, 2007). Low levels of androgens following castration increase pituitary GTHs via negative

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feedback. This increase in GTHs may act directly on seminal vesicle cells or indirectly via increased androgen production by the interrenals (Chowdhury & Joy, 2007). In contrast, in the long-jawed mudsucker (Gillichthys mirabilis, Gobiidae, Perciformes), castration leads to regression of the seminal vesicles, which is similar to what occurs in mammals (Chowdhury & Joy, 2007). These results indicate that seminal vesicles are steroidogenic and responsive to androgens and/or GTHs.

6.2.4. Components of seminal vesicle plasma and their functions 6.2.4.1. Ionic concentration, pH, and osmolality There are very few studies on seminal vesicle plasma (reviewed in Chowdhury & Joy, 2007). Although not specifically studied to date, we expect that hormones involved in whole-body osmoregulation are also important in controlling ionic concentration, pH, and osmolarity in seminal fluid. These hormones include cortisol, prolactin, GH, IGF-I, vasotocin, urotensins, and natriuretic peptides (Takei & Loretz, 2006). In the species that have been studied (the catfishes H. fossilis and C. gariepinus (Siluriformes) and the grass goby Zosterisessor ophiocephalus (Perciformes)), differences in the composition of seminal vesicle plasma relative to testicular and blood plasma have been documented (Chowdhury & Joy, 2007). Seminal vesicle plasma is colorless and highly viscous. It has a pH close to neutral and high specific gravity and osmolality relative to testicular or blood plasma. The major cations present in seminal vesicle plasma are Naþ, Kþ, and Ca2þ. Naþ concentration is comparable to that of blood plasma, but is higher than that of testicular plasma. Ca2þ and Kþ concentrations are higher in seminal vesicle plasma than in blood plasma, but are comparable to testicular plasma concentrations. Zn2þ, Cu2þ, and Mg2þ are also present, but at lower concentrations. Concentrations of these cations have been found to vary greatly among species. The pH, ionic concentration, and osmolality of seminal vesicle plasma all play a role in sperm motility (Chowdhury & Joy, 2007). Ionic gradient is important for the induction of sperm motility and for sperm longevity (Scaggiante et al., 1999; Rasotto & Mazzoldi, 2002; Mazzoldi, Scaggiante, Ambrosin, & Rasotto, 2000; Mazzoldi, Petersen, & Rasotto, 2005). Sperm motility in undiluted seminal vesicle plasma is suppressed due to higher ratios of Naþ and Ca2þ to Kþ. However, when seminal vesicle plasma is diluted (as occurs upon release into the water), the relative cation concentration decreases, neutralizing this motility suppression (Morisawa, Suzuki, & Morisawa, 1983; Mansour, Lahnsteiner, & Patzner, 2002; 2004; Alavi & Cosson, 2006; Morita et al., 2006). In the testis, low pH decreases sperm motility (Miura,


Yamauchi, Takahashi, & Nagahama, 1992). Higher secretions of bicarbonate ions into seminal vesicle plasma increase pH. Therefore, sperm exposed to this increase in pH have increased motility when released into the environment (e.g., Scott & Baynes, 1980; Baynes, Scott, & Dawson, 1981; Ohta, Ikeda, & Izawa, 1997; Wang & Crim, 1997). Osmotic pressure also plays a role in sperm motility, whether sperm are released into a more or less hyperosmotic environment (fresh or marine water). The high viscosity of seminal vesicle plasma slows the mixing of sperm with water, which protects them from the osmotic shock that would occur with rapid mixing. Osmotic shock can result in changes in membrane permeability and function, so preventing osmotic shock helps to maintain the mitochondrial integrity necessary for sperm motility and increases sperm longevity (Morisawa, Suzuki, & Morisawa, 1983; Stoss, 1983; Poupard et al., 1997; Alavi & Cosson, 2005; 2006). Some effects of seminal vesicle plasma on sperm activity are similar to what has been seen in response to exposure of sperm to ovarian fluids, which may also modify the behavior of sperm. For example, ovarian fluid increases sperm longevity, swimming speed, and linearity of movement in Arctic char (Salvelinus alpinus, Salmonidae, Salmoniformes) (Turner & Montgomerie, 2002). Mg2þ and Naþ in the ovarian fluid may increase motility and longevity of sperm (e.g., Linhart, Cosson, Mims, Shelton, & Rodina, 2002; Alavi, Cosson, Karami, Amiri, & Akhoundzadeh, 2004). Further discussion of ovarian fluid is obviously outside the scope of this review, but we wanted to alert the reader to the effects of this fluid on sperm. 6.2.4.2. Lipids, carbohydrates, and proteins There is currently no detailed analysis of classes of lipids present in seminal vesicle plasma. However, free and esterified cholesterol and phospholipids have been found. They are present at low concentrations, relative to concentrations seen in blood plasma (Mansour et al., 2004; Lahnsteiner & Patzner, 2009a). More detailed lipid analysis remains an important area for future investigations, especially those related to sperm performance. Carbohydrates are found in seminal vesicle plasma in the form of glucose, fructose, sucrose, and hexosamines, with concentrations varying among species (reviewed in Lahnsteiner & Patzner, 2009a). While in their immotile state, fish sperm obtain energy from endogenous respiration of intracellular lipids, glycolysis, tricarboxylic acid cycle, oxidative phosphorylation, and b-oxidation (Mansour, Lahnsteiner, & Berger, 2003). Seminal vesicles secrete mucopolysaccharides and sialomucins (reviewed in Chowdhury & Joy, 2007; Lahnsteiner & Patzner, 2009a). As mentioned above for testicular glands, these substances increase the viscosity of

Chapter | 3


seminal vesicle plasma, which aids in the agglutination of sperm and in some species helps adhere sperm to the substrate upon which females deposit eggs (e.g., Ota, Marchesan, & Ferrero, 1996; Scaggiante et al., 1999). Proteins are the main organic constituents of seminal vesicle plasma. They have been investigated via enzyme activity and electrophoresis in a few teleost species and can be classified as neutral, glycoproteins, or lipoproteins (e.g., Loir et al., 1990; reviewed in Chowdhury & Joy, 2007). However, the specific compositions are unknown. Seminal vesicle plasma also has a high concentration of lytic enzymes (including glucuronidase) and various protease inhibitors (e.g., Lahnsteiner et al., 1993; Mansour et al., 2004). Glycosidases and catabolic enzymes in the seminal vesicle plasma may function in both autophagic processes, involving mainly the degradation of secretory vesicles, and heterophagic processes, involving resorption of sperm (Lahnsteiner, Patzner, & Weismann, 1994; Chowdhury & Joy, 2007). 6.2.4.3. Steroid glucuronides and olfaction Males of many fish species release water-borne steroids and prostaglandins that have been shown to induce reproductive and behavioral responses in conspecifics (Van den Hurk & Resink, 1992; Stacey, Chojnacki, Narayanan, Cole, & Murphy, 2003; Scott et al., 2010) (see Chapter 7, this volume). During the breeding season, a broad spectrum of steroid glucuronides, primarily in the 5b-reduced form, occurs in the seminal vesicles of the African catfish C. gariepinus (Schoonen et al., 1988; Lambert & Resink, 1991). These steroid glucuronides may act as pheromones upon release into the environment (Van den Hurk et al., 1987; Lambert & Resink, 1991; Van den Hurk & Resink, 1992). In C. gariepinus and H. fossilis, glucuronated steroids secreted from the seminal vesicles act as olfactory stimuli for females, decreasing their brain DA content and increasing norepinephrine (NE) levels and LH secretion (Van den Hurk, Resink, & Peute, 1987; Lambert & Resink, 1991).

7. INTRASPECIFIC VARIATION IN SPERM CHARACTERISTICS AND TESTICULAR FUNCTION Species-typical patterns of gonadal recrudescence and regression across an annual cycle or breeding season are known for many fishes (e.g., Breder & Rosen, 1966; Munro et al., 1990). In addition to this annual variation in testicular structure and function, intraspecific variation also exists, often associated with variation in male reproductive tactics. In numerous fish species, there are discrete differences among males in the tactics used to deliver sperm. In these species, males practice what are referred to as alternative reproductive tactics (ARTs) (Oliveira, Cana´rio, & Ros, 2008). In species with ARTs, some males achieve matings

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by displaying to females and guarding access to them, the spawning sites, and/or the eggs and fry. Other males spawn using more surreptitious means, such as those used by males exhibiting ‘cuckolder’ tactics (e.g., sneaker, satellite, or female-mimicking males) (reviewed in Taborsky, 1994; Knapp & Neff, 2008; Oliveira et al., 2008; Taborsky, 2008). Sneaker males typically dart quickly towards a spawning pair to release sperm and then quickly retreat. Satellite or female-mimicking males, in contrast, rely on visual, and sometimes behavioral, mimicry of females to avoid detection by displaying males such that these males typically spawn in very close proximity to the female and rapid retreats are not necessary. Species with ARTs have been the focus of considerable study regarding behavioral aspects of reproduction (see Oliveira et al., 2008). Very few studies to date have focused on potential intraspecific variation in spermatogenesis among males exhibiting ARTs, much less the hormonal control thereof (see also Montgomerie & Fitzpatrick, 2009). Species with male ARTs should be particularly informative for studying both endocrine and other factors controlling spermatogenesis because the different male phenotypes are found to differ in some testicular and sperm features. Such differences likely have been selected for as a result of competition among sperm from different males, which is widespread within fishes and typifies those species with ARTs (Parker, 1990; Stockley, Gage, Parker, & Møller, 1997; Parker, 1998; Montgomerie & Fitzpatrick, 2009). Specifically, because cuckolder males experience a higher risk of sperm competition than males that guard eggs, cuckolders are expected to invest proportionately more into sperm number or quality (Parker, 1990); cuckolders typically cannot mate without a guarding male present, but guarders can and do spawn in the absence of cuckolder males. Relative investment in sperm is typically described with the GSI. Across fishes, GSI usually correlates positively with the intensity of sperm competition (Stockley et al., 1997; but see Pyron, 2000) and, within species, cuckolders usually have larger GSIs than guarding males (e.g., Gross, 1982; Taborsky, 1988; Montgomerie & Fitzpatrick, 2009). However, GSI by itself actually tells us very little about whether and how the sperm themselves might vary across male phenotypes, or even about the rate of spermatogenesis. Few studies have investigated more specific aspects of spermatogenesis and testicular function in fishes with ARTs. Here, we briefly summarize information about hormones and testicular function in three fish species with ARTs. The two male phenotypes in the protogynous saddleback wrasse Thalassoma duperrey (Labridae, Perciformes) differ in gonad size, histology, and steroid production (Hourigan, Nakamura, Nagahama, Yamauchi, & Grau, 1991). Large terminal phase (TP) males that have undergone sex change from female to male defend temporary

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spawning territories and have smaller testes than the smaller initial phase (IP) males that spawn in groups. Initial phase males undergo sexual maturation directly into a reproductive male, not passing first through a period of reproducing as a female. Testes of TP males have more numerous Leydig cells and produce more T and 11-KT in vitro. However, both testes types produce 17,20b-DHP at similar rates and at highest levels during the spawning period. Based on these results, the authors speculate that 17,20b-DHP is involved in spermiation but that 11-KT production is related more to male phenotype than to spermatogenesis. Neat, Locatello, and Rasotto (2003) studied reproductive morphology in male combtooth blennies that exhibit ARTs (Scartella cristata, Blenniidae, Perciformes). This species has three types of male: nesters, big males that care for eggs; hole-dwellers, medium-sized nonreproducing males that are site-attached to a hole; and sneakers, small males that release sperm in the nests of the big males. The anal fin gland is relatively larger in nesters than in holedwellers and is rudimentary in sneakers. The GSI of


sneakers is greater than the GSIs of nesters or holedwellers, but nesters and hole-dwellers exhibit a highly developed testicular gland. Nesters and hole-dwellers also possess a pair of secretory blind pouches that are barely visible in sneakers. Sperm duct walls of nesters are thickened and highly secretory, containing sperm dispersed in a granular matrix; in sneakers they are thin-walled and packed with concentrated sperm. Bluegill sunfish (L. macrochirus, Centrarchidae, Perciformes) are currently the best-studied species with respect to within-sex variation in testicular function. There are three alternative male tactics in the bluegill: large displaying parental males and two smaller cuckolder morphs (Gross, 1982). The cuckolder morphs (satellites and sneakers) are parasitic on the parental male’s care of the offspring (Neff, Fu, & Gross, 2003). In addition to behavioral differences, male morphs also differ in plasma steroid hormone concentrations and several sperm characteristics. On the day of spawning, parental males and cuckolders differ in plasma concentrations of T, 11-KT, E2, and cortisol (Figure 3.5) (Kindler, Philipp, Gross, & Bahr, 1989; Knapp

FIGURE 3.5 Variation in spermatogenesis and plasma hormone levels in the three male phenotypes of bluegill sunfish (Lepomis macrochirus, Perciformes, Centrarchidae) on the day of spawning. The male phenotypes experience different levels of sperm competition, which may have selected for variation in spermatogenesis; see text for description of mating behavior of each phenotype. The upper panels are photomicrographs of a testis from a representative male of each phenotype. Note the parental male testis contains more densely packed sperm in the lumen of the lobules compared with the relatively empty lumen of the satellite and sneaker males. Also note in the satellite and sneaker testes the greater proportion of cysts containing spermatocytes compared with the parental male testis, which contains more cysts with sperm ready to be released into the lumen. The lower left panel shows quantification of cysts containing the indicated stage of spermatogenesis (R. Knapp, B.D. Neff, & C.J. Leary, unpublished data). The lower right panel shows plasma levels of testosterone and 11-ketotestosterone (11-KT) (left axis) and cortisol (right axis) from these same males (data from Knapp & Neff, 2007); n ¼ 9e20 per male phenotype. SG-A, type A spermatogonia; SG-B, type B spermatogonia; SC, spermatocytes; ST, spermatids; SZ, sperm.

Chapter | 3

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& Neff, 2007). Across species with ARTs, 11-KT has been consistently associated with males that use a display tactic to attract females for spawning, but the role of T, if any, in behavioral tactic differences is unclear (Brantley, Wingfield, & Bass, 1993; Borg, 1994; Knapp, 2003; Oliveira, 2006). Cuckolders have smaller testes than parentals but invest proportionately more into testes as measured by GSI (Neff, Fu, & Gross, 2003). Sneakers and satellites also produce milt that contains higher concentrations of sperm than that of parentals (Stoltz & Neff, 2006). The sperm from sneakers contain more ATP, have slightly longer flagella, and swim faster than sperm from parentals (Neff et al., 2003; Burness, Casselman, SchulteHostedde, Moyes, & Montgomerie, 2004; but see Burness, Moyes, & Montgomerie, 2005). However, parental males’ sperm are longer-lived that those of cuckolders (Leach & Montgomerie, 2000; Neff et al., 2003). Interestingly, the latter trait differs from three other studied fish species with ARTs, where sneaker-type males have longer-lived sperm than nest guarders: three-spined stickleback (Gasterosteiformes) (De Fraipont, FitzGerald, & Guderley, 1993), Atlantic salmon (Salmoniformes) (Gage, Stockley, & Parker, 1995), and corkwing wrasse (Symphodus melops, Labridae, Perciformes) (Uglem, Galloway, Rosenqvist, & Folstad, 2001). The physiological mechanisms underlying these morph differences in bluegill sperm characteristics are currently unknown but have recently become a focus of our laboratory. To date, we have determined that male morphs differ in testis histology and pattern of spermatogenesis (Figure 3.5) (R. Knapp, B. D. Neff, & C. J. Leary, in preparation). Additional studies are underway to determine whether the male morphs differ in the timing of active spermatogenesis, testicular steroidogenic enzyme activity (especially 11b-HSD), and rate of spermatogenesis. Planned microarray studies may identify candidate genes contributing to individual variation in spermatogenesis, capitalizing on the fact that we can compare males with three different patterns of spermatogenesis within a single species. The sex steroids, via their role as transcription factors, may orchestrate molecular interactions that influence the different sperm characteristics documented in alternative male morphs (e.g., bluegills) or the various types of sperm (e.g., parasperm in sculpins described above) (see also Rolland et al., 2009).

8. CONCLUSIONS Understanding testicular structure and function in fishes is important for both basic and applied reasons. Fishes are a particularly powerful group for investigating endocrine signaling pathways of testicular function and spermatogenesis, and coordination with reproductive behavior. The cystic nature of spermatogenesis in fishes facilitates

molecular dissection of the various chemical messengers controlling particular stages of spermatogenesis. The diversity of reproductive modes in fishes provides a rich resource for comparative studies of endocrine control of testicular function. The genome-wide duplication that occurred at the appearance of the Teleostei has provided many genes that could be explored for studying protein function as related to relatively small changes in structure and in comparison to the function of the single copy of these genes in the non-teleost fishes. Finally, advances in our understanding of fish phylogeny, together with the increasing ease of sequencing and other molecular techniques, now makes more feasible the addressing of very challenging questions regarding testicular function, even in ‘non-model’ species. An excellent example of this is the recent elucidation of the timing of AR gene duplication in fishes and the proposed neofunctionalization of the ARb gene (i.e., acquisition of new function(s) relative to the ARa gene) in percomorphs (Douard et al., 2008). By noting an intriguing coincidence of the origin of the proposed neofunctionalization of the ARb gene and the occurrence of viviparity given our current understanding of fish phylogeny, we propose that ARb will be an important molecule to investigate for understanding the evolution of viviparity in the Acanthopterygii. Similarly, comparing the role of AR in testis function in non-teleost fishes with the role of the single or duplicated AR genes in teleosts is yet another fruitful area for future research. The coming years promise true exploration and exploitation of the great diversity of fishes in order to understand the evolution of hormonal regulation of testicular and accessory structure function, as well as the mechanisms that underlie coordination of those functions with variation in male reproductive behavior.

ACKNOWLEDGMENTS We sincerely thank David Norris and Kristin Lopez for the invitation to contribute this chapter and for their patience, and Pat Gonzalez at Elsevier for her assistance with production. We also thank Richard Broughton (Principle Investigator for the Euteleost Tree of Life project funded by the National Science Foundation) for helping to ensure our representation of fish phylogeny in Figure 3.3 reflects current consensus based on the various datasets currently available. Finally, we thank John Lukeman for organizing the data that are summarized in Figure 3.3 and Tamam Al-Ali for assistance with the final stages of manuscript preparation. Our work on sunfish was supported by the National Science Foundation (IBN/IOS 0349449) to R. Knapp and carried out in collaboration with Bryan Neff.

ABBREVIATIONS 11b-HSD 11-KT 17,20bDHP

11b-hydroxysteroid dehydrogenase 11-ketotestosterone 17,20b-dihydroxypregn-4-en-3-one

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17,20aDHP AND AR ART DA E2 ELISA FSH GH GnRH GSI GTH HPLC IGF-I IGF-II IP Kp LC-MS LH MEL NE OHA OHT P450C17 PGF2a RIA SG-A SG-B sGnRH SSC StAR T TGFb TP


17,20a-dihydroxypregn-4-en-3-one Androstenedione Androgen receptor Alternative reproductive tactic Dopamine Estradiol Enzyme-linked immunosorbent assay Follicle-stimulating hormone Growth hormone Gonadotropin-releasing hormone Gonadosomatic index Gonadotropin High pressure liquid chromatography Insulin-like growth factor-I Insulin-like growth factor-II Initial phase Kisspeptin Liquid chromatography-mass spectrometry Luteinizing hormone Melatonin Norepinephrine 11b-hydroxyandrostenedione 11b-hydroxytestosterone 17a-hydroxylase/17,20b-lyase Prostaglandin F2a Radioimmunoassay Type A (primary) spermatogonia Type B (secondary) spermatogonia Salmon gonadotropin-releasing hormone Spermatogonial stem cells Steroidogenic acute regulatory Testosterone Transforming growth factor-b Terminal phase

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Schoonen, W., & Lambert, J. G. D. (1986). Steroid-metabolism in the seminal vesicles of the African catfish, Clarias gariepinus (Burchell), during the spawning season, under natural conditions, and kept in ponds. Gen. Comp. Endocrinol., 61, 355e367. Schoonen, W., Lambert, J. G. D., & Vanoordt, P. (1988). Quantitative analysis of steroids and steroid glucuronides in the seminal vesicle fluid of feral spawning and feral and cultivated nonspawning African catfish, Clarias gariepinus. Gen. Comp. Endocrinol., 70, 91e100. Schlupp, I. (2005). The evolutionary ecology of gynogenesis. Annu. Rev. Ecol. Syst., 36, 399e417. Schulz, R. W., & Miura, T. (2002). Spermatogenesis and its endocrine regulation. Fish Physiol. Biochem., 26, 43e56. Schulz, R. W., Menting, S., Bogerd, J., França, L. R., Vilela, D. A. R., & Godinho, H. P. (2005). Sertoli cell proliferation in the adult testis: evidence from two fish species belonging to different orders. Biol. Reprod., 73, 891e898. Schulz, R. W., De França, L. R., Lareyre, J.-J., LeGac, F., ChiariniGarcia, H., No´brega, R. H., et al. (2010). Spermatogenesis in fish. Gen. Comp. Endocrinol., 165, 390e411. Scott, A. P., & Baynes, S. M. (1980). A review of the biology, handling and storage of salmonid spermatozoa. J. Fish Biol., 17, 707e739. Scott, A. P., Sumpter, J. P., & Stacey, N. (2010). The role of the maturation-inducing steroid, 17,20 b-dihydroxypregn-4-en-3-one, in male fishes: a review. J. Fish Biol., 76, 183e224. Seiwald, M., & Patzner, R. A. (1989). Histological, fine-structureal and histochemical differences in the testicular glands of gobiid and blenniid fishes. J. Fish Biol., 35, 631e640. Senthilkumaran, B., & Joy, K. P. (1993). Annual cyclic, and castration and cyproterone acetate-induced, changes in sialic acid content of the seminal vesicle of the catfish, Heteropneustes fossilis (Bloch). Fish Physiol. Biochem., 10, 425e430. Singh, M. S., & Joy, K. P. (1998). A comparative study on histochemical distribution of some enzymes related to steroid and glucuronide synthesis in seminal vesicle and testis of the catfish. Clarias batrachus. Zool. Sci., 15, 955e961. Singh, M. S., & Joy, K. P. (1999). Annual correlative changes in some biochemical contents of seminal vesicle and testis in the catfish Clarias batrachus (L). Zool. Sci., 16, 345e356. Small, C. M., Carney, G. E., Mo, Q., Vannucci, M., & Jones, A. G. (2009). A microarray analysis of sex- and gonad-biased gene expression in the zebrafish: Evidence for masculinization of the transcriptome. BMC Genomics, 10, 579, doi:10.1186/1471-2164-10-579. Sorensen, P. W., Pinillos, M., & Scott, A. P. (2005). Sexually mature male goldfish release large quantities of androstenedione into the water where it functions as a pheromone. Gen. Comp. Endocrinol., 140, 164e175. Sperry, T. S., & Thomas, P. (1999). Characterization of two nuclear androgen receptors in Atlantic croaker: Comparison of their biochemical properties and binding specificities. Endocrinology, 40, 1602e1611. Sreenivasan, R., Cai, M., Bartfai, R., Wang, X., Christoffels, A., & Orban, L. (2008). Transcriptomic analyses reveal novel genes with sexually dimorphic expression in the zebrafish gonad and brain. PLoS One, 3, e1791. Stacey, N., Fraser, E., Sorenson, P., & Van Der Kraak, G. (2001). Milt production in goldfish: regulation by multiple social stimuli. Comp. Biochem. Physiol. C, 130, 467e476.

Chapter | 3


Stacey, N., Chojnacki, A., Narayanan, A., Cole, T., & Murphy, C. (2003). Hormonally derived sex pheromones in fish: exogenous cues and signals from gonad to brain. Can. J. Physiol. Pharmacol., 81, 329e341. Stiassny, M. L. J., Wiley, E. O., Johnson, G. D., & De Carvalho, M. R. (2004). Gnathostome fishes. In J. Cracraft, & M. J. Donoghue (Eds.), Assembling the tree of life (pp. 410e429). New York, NY: Oxford University Press. Stockley, P., Gage, M. J. G., Parker, G. A., & Møller, A. P. (1997). Sperm competition in fishes: the evolution of testis size and ejaculate characteristics. Am. Nat, 149, 933e954. Stoltz, J. A., & Neff, B. D. (2006). Sperm competition in a fish with external fertilization: the contribution of sperm number, speed, and length. J. Evol. Biol., 19, 1873e1881. Stoss, J. (1983). Fish gamete preservation and spermatozoan physiology. Fish Physiol., 9, 305e350. Sundararaj, B. I., & Nayyar, S. K. (1969a). Effects of castration and/or hypophysectomy on seminal vesicles of catfish, Heteropneustes fossilis (Bloch). J. Exp. Zool., 172, 369e384. Sundararaj, B. I., & Nayyar, S. K. (1969b). Effects of estrogen, SU-9055, and cyproterone acetate on hypersecretory activity in seminal vesicles of castrate catfish, Heteropneustes fossilis (Bloch). J. Exp. Zool., 172, 399e408. Taborsky, M. (1994). Sneakers, satellites, and helpers: Parasitic and cooperative behavior in fish reproduction. Adv. Study Behavior., 23, 1e100. Taborsky, M. (1998). Sperm competition in fish: ‘bourgeois’ males and parasitic spawning. Trends Ecol. Evol., 13, 222e227. Taborsky, M. (2008). Alternative reproductive tactics in fish. In R. F. Oliveira, M. Taborsky, & H. J. Brockmann (Eds.), Alternative reproductive tactics (pp. 251e299). Cambridge, UK: Cambridge University Press. Takei, Y., & Loretz, C. A. (2006). Endocrinology. In D. H. Evans, & J. B. Claiborne (Eds.), The physiology of fishes (3rd Ed., pp. 271e318). Boca Raton, FL: CRC Press. Takemura, A., Rahman, M. S., Nakamura, S., Park, Y. J., & Takano, K. (2004). Lunar cycles and reproductive activity in reef fishes with particular attention to rabbitfishes. Fish and Fisheries, 5, 317e328. Takeo, J., & Yamashita, S. (1999). Two distinct isoforms of cDNA encoding rainbow trout androgen receptors. J. Biol. Chem., 274, 5674e5680. Tanaka, M., Kinoshita, M., Kobayashi, D., & Nagahama, Y. (2001). Establishment of medaka (Oryzias latipes) transgenic lines with the expression of green fluorescent protein fluorescence exclusively in germ cells: a useful model to monitor germ cells in a live vertebrate. Proc. Natl. Acad. Sci. USA, 98, 2544e2549. Thacker, C., & Grier, H. (2005). Unusual gonad structure in the paedomorphic teleost Schindleria (Teleostei: Gobioidei), with a comparison to other gobioid fishes. J. Fish Biol., 66, 378e391. Turner, C. L. (1960). The effects of steroid hormones on the development of some secondary sexual characters in cyprinodont fishes. Trans. Am. Microsc. Soc., 79, 320e333. Turner, E., & Montgomerie, R. (2002). Ovarian fluid enhances sperm movement in Arctic charr. J. Fish Biol., 60, 1570e1579. Uglem, I., Galloway, T. F., Rosenqvist, G., & Folstad, I. (2001). Male dimorphism, sperm traits and immunology in the corkwing wrasse (Symphodus melops L.). Behav. Ecol. Sociobiol., 50, 511e518. Uribe Aranza´bal, M. C., Grier, H. J., De la Rosa Cruz, G., & Garcıá Alarcoń, A. (2009). Phylogeny and classification. In B. G. M. Jamieson (Ed.), Reproductive biology and phylogeny of

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Chapter 4

Regulation of Ovarian Development and Function in Teleosts R. Urbatzka,* M.J. Rocha*, y and E. Rocha* *

University of Porto, Porto, Portugal, y Superior Institute of Health Sciences North (ISCS-N), Paredes, Portugal

SUMMARY The genesis and development of oocytes in teleost fishes is a fascinating process with tight and complex physiological regulation mechanisms that ensure the organization of genetic and nutritional bases for the development of new individuals. Female gametogenesis comprises several developmental steps and consists of oogenesis, oocyte growth, maturation, and ovulation. Primordial germ cells differentiate into oogonia under the influence of early gene cascades and steroid signaling. Oocyte growth is triggered by gonadotropins, and the subsequent steroidogenic production of estradiol induces vitellogenesis, leading to a marked enlargement in oocyte size. Neuroendocrine factors including stimulatory and inhibitory signals are regarded as major regulators of oocyte development and mediators of environmental and physiological cues. Maturation requires meiotic resumption, and is triggered both by gonadotropins and maturation-inducing hormone. Gonadotropin signals are mediated or modulated in the ovary by a complex local paracrine network of peptide factors.

1. INTRODUCTION: FISH MODELS OF REPRODUCTIVE STRATEGIES Fish reproduction is characterized by a puzzling diversity of strategies regarding how different species determine their sex (see Chapter 1, this volume), produce male and female fertile gametes, and mate. Maybe the success of the teleost group in conquering so many different habitats around the world is owed, at least in part, to these special features. Sex in fishes may be determined genetically, by environmental factors, or by social behavior (see also Chapter 8, this volume). Temperature is an important environmental factor, and an influence on sex determination is observed in several groups of fishes. The effect of temperature is seen at a critical sensitive period of development, and lower temperatures increase the male

Hormones and Reproduction of Vertebrates, Volume 1dFishes Copyright Ó 2011 Elsevier Inc. All rights reserved.

to female ratio or promote ovarian differentiation (Baroiller & D’Cotta, 2001). A prominent example of sex determination by social behavior is the clownfish (Amphiprion percula) inhabiting anemones, where mating groups have a dominant female, a mating male, and several other males. If the dominant female dies, the mating male turns into a female, and one of the previously nonmating males turns into the mating male (Buston, 2004). Species may be gonochoristic or hermaphroditic, the latter mostly separable into protandrous or protogynous, as reviewed in Rocha and Rocha (2006). In gonochoristic species, male and female sex is always clearly distinct (not always phenotypically) and maintained over time. Hermaphroditism in teleosts is usually sequential, meaning that in most species individuals develop first as one sex and later transform into the opposite sex. In protandrous hermaphroditic species, the immature fish develop first as males and later as females, whereas in protogynous species the opposite situation exists. The sharpsnout seabream (Diplodus puntazzo) is an example of a species that exhibits protandry (Pajuelo, Lorenzo, & Dominguez-Seoane, 2008), whereas protogyny, the most common form of hermaphroditism, is often described in wrasses (Labridae) (Morton, Gladstone, Hughes, & Stewart, 2008) and groupers (Serranidae) (Liu & De Mitcheson, 2009). In some sequential hermaphrodites, all immature fish exhibit male or female development and a distinct percentage (usually of one sex) proceed further to the opposite sex. In contrast, simultaneous hermaphrodites (that display both male and female gonadal features) are rare in fishes, but this does occur in some serranids (Leonard, 1993; Brauer, Scharer, & Michiels, 2007). Many differences exist in the maturation strategy of gametes in diverse fish species (see Rocha & Rocha, 2006). Synchronous spawners mature gametes in both sexes simultaneously and spawn all at the same time. Often these

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individuals die after spawning; Pacific salmon (Oncorhynchus spp.) and sea lampreys (Petromyzon marinus) are examples of such semelparous groups. Group-synchronous spawners possess clusters of gametes in different stages and spawn during the course of a breeding season. Two stages of oocytes are the dominant populations of gametes. This is the most common mode of spawning in fishes, even if the specific time of breeding may be different. Examples include rainbow trout (Oncorhynchus mykiss), Atlantic salmon (Salmo salar), and European sea bass (Dicentrarchus labrax). Asynchronous spawners have the potential to spawn during the whole year and all stages of gametes are present in the gonads of mature fish at all times. No dominant developmental stage is observed within the oocyte populations. Many tropical fishes belong to this group and common examples are tilapia (Oreochromis spp.), killifishes (Fundulus spp.), and two model fish species used in laboratories around the world: zebrafish (Danio rerio) and medaka (Oryzias latipes) (Ankley & Johnson, 2004). The aforementioned diversity of reproductive strategies in fishes is in sharp contrast to the reproductive scheme of mammals, birds, and many reptiles and amphibians, generally characterized by genetically sex-determined gonochorism. However, the fundamental processes underlying gametogenesis and their physiological regulation are similar among vertebrates, reflecting the evolutionary importance of these vital processes. Fishes have classically been good models for studying the reproductive biology of vertebrates and, from a comparative view, the endocrine and paracrine regulation of reproduction in fishes underlying different reproductive strategies offers great potential to obtain further insights into reproduction in all vertebrates. Fishes are easy to use in experiments and to manipulate, often having short generation times or eggs that are transparent, allowing the study of early development. Further, genomic information is becoming more and more available. None-the-less, progress in the field of fish reproduction is mainly restricted to a few species, namely those with commercial interest or those frequently used as model species for a range of purposes; e.g., rainbow trout and other salmonids, medaka and zebrafish, red seabream (Pagrus major), European sea bass, and eel (Anguilla anguilla). In short, the diversity of reproductive strategies of fishes offers a great potential to gain further insights into the process of fish/vertebrate gametogenesis and its regulation by endocrine and paracrine factors, and the roles of nutrition, body growth, and environmental factors. Additionally, fundamental knowledge about the development of fertilizable gametes in fishes and their physiological regulation will be useful for the research fields of aquaculture, reproductive toxicology, and biomedical applications.


2. MORPHOLOGICAL ASPECTS OF THE TELEOST OVARY AND STAGES OF OOCYTE DEVELOPMENT The basic events of oocyte development apply to most fishes. Primordial germ cells (PGCs) develop into oogonia that proliferate, thus increasing in number, and subsequently develop into primary growth oocytes (PGOs). Primary growth oocytes are enclosed by follicular cells, grow while arrested in the meiotic division of prophase I, and later increase dramatically in size due to hepatic vitellogenesis and subsequent incorporation of vitellogenin (Vtg). For maturation, the grown oocytes resume meiosis, which can be visibly observed as the peripheral migration of the nucleus, the germinal vesicle (GV), and later as GV breakdown (GVBD). Fertilizable oocytes usually are then ovulated into the ovarian lumen and soon after expelled via gonoducts for external fertilization. However, a few fish species fertilize their oocytes while within the ovary, as in guppies and mollies (Poeciliidae: Poeciliinae). Departures from the common model of ovarian anatomy and ovulation exist, as in salmonids, where the ovarian lumen is continuous with the coelom dorsally, and mature oocytes are ovulated first into the abdominal cavity, where they can remain for over a week before being truly spawned via the oviduct for fertilization by waiting males (Springate, Bromage, Elliott, & Hudson, 1984). When oocytes are released directly into the coelomic cavity, the ovary is said to be of the gymnovarian type (most teleosts) as opposed to the cystovarian type (e.g., guppies). Irrespective of the type, the ovary in most teleosts is a paired organ that is attached to the body cavity. As mentioned, oocytes generally are ovulated into the ovarian cavity (intraovarian space), and oocytes are excreted via paired gonoducts that typically fuse before reaching the genital pore (Patino & Redding, 2000). The ovary normally consists of germ cells, oogonia, different stages of oocytes, granulosa cells, theca cells, stroma cells and connective tissue matrix, blood vessels, and nervous tissue. The development of oocytes can be divided into five stages (Figure 4.1), which are similar among the diverse teleost species (Selman, Wallace, Sarka, & Qi, 1993; Leino, Jensen, & Ankley, 2005). The sequence of oocyte stages is as follows: (I) primary growth, (II) cortical alveoli stage, (III) early vitellogenic stage, (IV) late vitellogenic stage, and (V) matured/ovulated. Stage I comprises PGOs, with small cytoplasmic volume, that are organized in cell nests together with oogonia and prefollicular cells (stage Ia). Subsequently, the PGOs enlarge, become surrounded by follicle cells, and leave the cell nests as primary follicles, the complex of oocytes with their somatic cells (stage Ib). The layer of somatic tissues already consists of granulosa cells (inner layer) and theca cells (outer layer). Both cell

Chapter | 4

Regulation of Ovarian Development and Function in Teleosts

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FIGURE 4.1 General scheme of oocyte development and growth in fishes. The sequence of oocyte stages is as follows: stage Idprimary growth; stage IIdcortical alveoli growth period; stage IIIdearly vitellogenic oocytes; stage IVdlate vitellogenic phase; and stage Vdmature/ovulated oocyte, full of yolk (with lipid and protein globules). Oocyte growth is controlled by 17b-estradiol (E2) and follicle-stimulating hormone (FSH), whereas the resumption of meiosis is regulated by luteinizing hormone (LH) and maturation-inducing hormone (MIH). GVBD, germinal vesicle breakdown. See color plate section.

layers are involved in steroidogenesis, the formation of sex steroids (Nagahama, 1994). Stage Ib oocytes possess many nucleoli at the border of the nuclear envelope, and the PGOs arrest in prophase I of meiosis. Stage Ib can be further characterized by two subcategories, the early and the late perinucleolus stages, according to oocyte size and cytoplasmic tinctorial affinity in histological sections, with increasing basophilic staining (Rocha & Rocha, 2006). Stage II of oocyte development is called the cortical alveolus stage because of the appearance of so-called cortical alveoli, defined as membrane-limited glycoprotein vesicles of variable size. (They are at times described in the literature as yolk vesicles, primary yolk, or endogenous yolk, but none of these names are particularly adequate in describing the alveolar content.) The cortical alveoli are synthesized for later participation in the cortical reaction that takes place after fertilization as a block against polyspermy. During the cortical reaction, the cortical alveoli are discharged into the perivitelline space between the oocyte and the vitelline envelope (zona radiata or chorion) (Patino, Yoshizaki, Thomas, & Kagawa, 2001; Patino & Sullivan, 2002). The vitelline envelope is essentially formed at stage II (it starts to form at the end of the primary growth stage), and is perforated by pore canals containing microvilli of

both the oocyte and follicle cells. The cortical alveolus stage is also termed the secondary growth stage (Rocha & Rocha, 2006). Stage III is the vitellogenic stage of the oocytes and is characterized as the major growth stage forming the enlarged tertiary follicle. Vitellogenin produced by the liver under the influence of estrogens secreted by the ovary is sequestered by the oocytes via endocytosis and then processed into yolk proteins. Cortical alveoli move to the periphery as yolk accumulates centripetally. Oocytes not only increase markedly in size, but the number of theca cells in the outer somatic cell layer also increases. Stage IV is the oocyte maturation period during which meiosis is reinitiated. The GV (enlarged oocyte nucleus arrested in prophase of meiosis I) migrates toward the periphery and breaks down during the first meiotic division (GVBD). Oocytes continue meiosis until the second meiotic metaphase, in which they are arrested until fertilization. During oocyte maturation in teleosts, many oocytes enlarge again in size due to hydration, which is particularly critical in marine species with pelagic eggs to ensure specific buoyancy. In many teleosts, oocytes become translucent during oocyte maturation due to disassembly of crystalline yolk proteins and subsequent fusion of yolk

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globules (Carnevali, Cionna, Tosti, Lubzens, & Maradonna, 2006). Stage V is the mature egg stage in that prior to ovulation the oocytes detach from the follicle cells. Mature eggs are ovulated into the ovarian lumen and are evacuated through the gonoduct in most species, while the remaining follicle cells are transformed into the postovulatory follicle. The completion of the second meiotic division of the mature egg occurs only after fertilization and results in the formation of a haploid egg and a polar body, with the haploid egg fusing with the haploid sperm. Fishes may display lifetime determined fecundity, annual determined fecundity, or indeterminate fecundity. In lifetime determined fecundity, the number of producible oocytes is established during early life and no further oocytes can be produced. In annual determined fecundity, the number of newly produced oocytes is determined during the beginning of each breeding season. Only in indeterminate fecundity is the number of producible oocytes not fixed, and oogonia are renewed continuously. In rainbow trout, a recent work strengthened the hypothesis of an active germinal epithelium that may give rise to new germ cells (Grier, Uribe, & Parenti, 2007). Such female teleosts may produce an indeterminate number of new oocytes from the germinal epithelium in each breeding season, although the number of spawned oocytes depends on a balance between folliculogenesis and oocyte atresia (Grier et al., 2007). The germinal epithelium is composed of both oogonia and somatic cells, and is separated from the stroma by a basement membrane. The somatic cells are epithelial cells able to become prefollicular cells if associated with a meiotic oocyte in the germinal epithelium. Together, prefollicular cells and oocytes initiate the formation of new follicles. The presence of an active germinal epithelium, the formation of cell nests, and ongoing folliculogenesis are important in the regulation of group-synchronous spawning. In contrast to many fish as well as amphibian and reptilian species with a renewable oogonial pool, fecundity in mammals and birds has been thought to be determined in the ovary around birth with a fixed number of oocytes that are arrested in prophase I of meiosis. However, as recently as in 2004, an active germinal epithelium was demonstrated in the postnatal mouse ovary, together with the production of new follicles (Johnson, Canning, Kaneko, Pru, & Tilly, 2004) that revolutionized this axiom of vertebrate reproductive biology. These findings, however, have not been replicated. Further, there is evidence and controversy supporting the formation of new primary follicles even in adult human ovaries (Bukovsky, Caudle, Svetlikova, & Upadhyaya, 2004; Bukovsky, Virant-Klun, Svetlikova, & Willson, 2006).


3. DIFFERENTIATION OF PRIMORDIAL GERM CELLS INTO OOGONIA An important step towards the development of a mature ovary is the early differentiation of primordial germ cells (PGCs) into oogonia, an early step in sexual differentiation. The gonadal anlagen at this early stage are bipotential and can be triggered by appropriate sex-specific gene cascades and steroid signaling pathways into male or female gonads (Hughes, 2001; Brennan & Capel, 2004). In mammals, the SRY gene located on the Y chromosome is responsible for genetic sex determination. If the SRY gene is expressed in the gonads, a gene cascade is initiated, including SOX9 and DMRT1, which trigger the development of a testis whereas otherwise an ovary would arise (Hughes, 2001; Brennan & Capel, 2004). The testis developmental pathway involves the subsequent emergence of Sertoli and Leydig cells. Sertoli cells secrete the anti-Mu¨llerian hormone (AMH), thus inducing regression of the Mu¨llerian ducts, and promote the emergence of the Leydig cells that secrete testosterone (T), responsible for both the development and maintenance of male secondary sex characteristics (Nef & Parada, 2000). If the ovarian developmental pathway is triggered by the absence of the SRY gene, a different set of genes is expressed including follistatin, wnt-4, and dax-1. Subsequently, both theca and granulosa cells develop, the latter secreting mainly 17b-estradiol (E2), thus ultimately leading to the development of ovaries (Hughes, 2001; Ross & Capel, 2005). In contrast to mammals, sexual differentiation in fishes, amphibians, and reptiles is more dynamic and regulated by genetic, endocrine, and environmental factors, as wellillustrated for fishes by the sea bass (D. labrax) (Piferrer, Blazquez, Navarro, & Gonzalez, 2005). Some of the genes involved in sexual differentiation have been discovered in lower vertebrates; however, the complex processes of early regulation of sexual differentiation are only partly understood and not all players and their complex interplay have been identified yet (Figure 4.2). In medaka, dmrt1 was identified as a major regulator of sexual differentiation comparable to the mammalian SRY gene (Nanda et al., 2002; Hornung, Herpin, & Schartl, 2007). Recent data from Nile tilapia confirmed that dmrt1 has a major role in testicular differentiation in Nile tilapia (Oreochromis niloticus) (Kobayashi, Kajiura-Kobayashi, Guan, & Nagahama, 2008). However, it may not be the universal sex-determining gene in teleosts since phylogenetic analyses showed that it was derived by an evolutionarily early (in fishes) gene duplication event (Volff, Kondo, & Schartl, 2003). Foxl2, follistatin, and ovarian aromatase (cyp19a1) show sexually dimorphic expression in rainbow trout, with higher expression in females suggesting a role in ovarian differentiation (Vizziano, Randuineau, Baron, Cauty, & Guiguen, 2007), as also shown in amphibians for cyp19a1

Chapter | 4


FIGURE 4.2 Overview of gonadal differentiation in fishes. Bipotential germ cells are triggered by sex-determining gene(s) into the male or female development pathway. Expression of sex-specific gene cascades and the occurrence of sex-specific steroidogenic cells and elevated steroid levels are important steps on the way to develop into males and females. It is noteworthy that in fishes no universal sex-determining gene has been identified, and that dmrt1 is the only identified gene, in medaka, with this particular function. Further, many deviations from this simplified scheme are present in fishes, including the impact of environmental factors (e.g., temperature) or of social behavior on sexual determination, and on protandrous and protogynous hermaphroditism.

(Urbatzka, Lutz, & Kloas, 2007). A similar result was obtained from the gonochoristic Nile tilapia (O. niloticus), in that foxl2 and cyp19a1 were only expressed in XX gonads during early gonadal differentiation from a screen of 17 candidate genes potentially involved in sexual differentiation (Ijiri et al., 2008). Restriction of aromatase gene expression to the ovary highlights the important roles of sexual steroids during gonadal differentiation. Both in rainbow trout and in Nile tilapia, aromatase mRNA expression was greater in females, but no differences were found for estrogen receptor (ER) mRNA expression (Guiguen et al., 1999). Aromatase gene expression and enzyme activity is regarded as one important trigger for gonadal differentiation, being repressed in testes and stimulated in ovaries (Guiguen, Fostier, Piferrer, & Chang, 2009). The gene foxl2 may be involved in the transcriptional regulation of cyp19a1 during early sexual differentiation (Ijiri et al., 2008). In vitellogenic follicles of Nile tilapia, Ad4BP (also known as steroidogenic factor (SF-1)) transcriptionally regulates cyp19a1, an indication of gonadotropin (GTH) regulation via second messenger(s) (Yoshiura et al., 2003). However, the expression patterns measured in ovaries during sex differentiation do not indicate cyp19a1 regulation via Ad4BP, but suggest transcriptional regulation by foxl2 (Ijiri et al., 2008). The link between foxl2 and cyp19a1 was first established in rainbow trout during normal gonadal differentiation as well as under feminizing or masculinizing conditions

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(Baron et al., 2004). During natural gonadal differentiation, the paralogs foxl2a and foxl2b are specifically expressed in the ovary and the expression profile of foxl2a is highly correlated to that of cyp19a1. Following 17a-ethinylestradiol treatment used to feminize genetically male trout, both paralogs were upregulated but cyp19a1 was not. In contrast, following 11b-hydroxyandrostenedione treatment or treatment with the aromatase inhibitor androstenetrione, both foxl2 transcripts were downregulated in females, again correlated to a downregulation of cyp19a1. The paralogs of foxl2 were studied in rainbow trout and sequence comparisons with several fish and mammalian species showed a high degree of similarity, thus indicating an evolutionary conservation from fishes to mammals. A role for foxl2a was suggested for ovarian differentiation, while foxl2b could be involved during the onset of first oocyte meiosis (Baron et al., 2004). The important role of sex steroids during gonadal differentiation has been impressively shown in animals exposed to exogenous sex steroids, which lead to sex reversal in fishes (Gimeno, Komen, Gerritsen, & Bowmer, 1998; Orn, Holbech, Madsen, Norrgren, & Petersen, 2003), amphibians (Bo¨gi, Levy, Lutz, & Kloas, 2002; Kloas, 2002), and reptiles (Crews, Bull, & Wibbels, 1991). In amphibians, exogenous steroids interfere with the hypothalamicepituitaryegonadal (HPG) axis (Urbatzka, Lutz, Opitz, & Kloas, 2006), gene expression patterns, and endogenous steroid levels (Urbatzka, Bottero, Mandich, Lutz, & Kloas et al., 2007), and alter gonadal structure and induce intersex gonads (Cevasco et al., 2008). The inhibition of endogenous E2 biosynthesis by inhibition of aromatase caused masculinization in the Japanese flounder (Paralichthys olivaceus) (Kitano, Takamune, Nagahama, & Abe, 2000) and in amphibians (Yu, Hsu, Ku, Chang, & Liu, 1993; Chardard & Dournon, 1999), and emphasized the special role of E2 in ovarian differentiation. These aforementioned studies demonstrated the potential of sex steroids to influence gonadal differentiation among lower vertebrates and the susceptibility of gonads to steroid signaling. However, there is little information on which stage(s) of gonadal development, and particularly on which developmental stage(s) of gametes, sexual steroids exert their impact to reverse normal gonadal differentiation. Germ cells have long been considered to be influenced by their surrounding somatic cell layers but not to play an active role in determining the fate of somatic cells. However, following ablation of germ cells in female mice, pregranulosa cells developed into immature Sertoli cells (Guigon, Coudouel, Mazaud-Guittot, Forest, & Magre, 2005). It was speculated that the complete differentiation of granulosa cells into Sertoli cells involves yet-unidentified factors and that the ability of follicle cells to switch to their male counterpart may reflect their similar fetal origin.

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Corresponding results are being obtained from studies in fishes. Recent evidence from medaka indicates that germ cells feminize gonadal somatic cells in the absence of masculinizing signals, and that gonadal somatic cells in the absence of germ cells are predisposed to a male developmental path (Kurokawa et al., 2007; Tanaka, Saito, Morinaga, & Kurokawa, 2008). Therefore, female germ cells are not only driven by signals emitted by their follicle cells, but are also involved in the determination of the fate of follicle cells. Consequently, germ cells may be important for the development of sexual dimorphism in fishes. This hypothesis would fit very well with the development of juvenile hermaphroditism in zebrafish, where, after initial development of all individuals as females, the loss of oocytes via atresia, namely by apoptosis (Uchida, Yamashita, Kitano, & Iguchi, 2002), could trigger the development of male somatic cells and subsequently the emergence of male gametes. In line with this hypothesis, the timing of this female-to-male transformation is variable in zebrafish (Wang, Bartfai, Sleptsova-Freidrich, & Orban, 2007) and the transformation is characterized by a decreased level of cyp19a and an increased level of AMH (Wang & Orban, 2007). It was reasoned that this supported an earlier differentiation of Sertoli cells than Leydig cells in the developing testis, while a tissular scenario of inhibition of Cyp19a1 could leave more substrates for Cyp11b (it uses the same substratedT as Cyp19a1) to produce 11-ketotestosterone (11-KT), increasingly promoting testicular growth.

4. OOGENESIS, OOCYTE GROWTH, AND DEVELOPMENT In vertebrates, oocyte growth and development are under the strict control of the HPG axis that regulates, via the secretion of GTHs, important processes such as follicular growth and steroidogenesis. In fishes, follicle-stimulating hormone (FSH) is the main GTH responsible for inducing growth and development of oocytes (Planas & Swanson, 2008). For example, during early gonadal differentiation of rainbow trout, FSH is measurable but luteinizing hormone (LH) is undetectable, thus suggesting a growth-promoting role for FSH in this period (Feist & Schreck, 1996). During further maturation and vitellogenesis in rainbow trout, FSH steadily increases whereas LH is increased during the maturational phase and spawning (Sumpter & Scott, 1989; Prat, Sumpter, & Tyler, 1996). Similar results were obtained from the European eel (A. anguilla), where FSH is high in immature females and decreases during maturation, whereas LH is initially low and increases during maturation and ovulation (Schmitz et al., 2005). These results described in salmonids (group-synchronous spawners) and for eels (lifetime synchronous spawners) are different from


the expression profiles of GTHs found in teleosts displaying asynchronous spawning. In goldfish (Carassius auratus), expression of FSH and LH mRNA steadily increased in parallel during asynchronous gametogenesis (Sohn, Yoshiura, Kobayashi, & Aida, 1999). In asynchronous spawning stickleback (Gasterosteus aculeatus), FSH mRNA expression precedes that of LH, which was associated with T levels and gonadosomatic index (GSI) (Hellqvist, Schmitz, Mayer, & Borg, 2006). In the red seabream (P. major), LH increased during vitellogenesis and decreased thereafter, whereas FSH was increased at spawning and decreased from then on (Gen et al., 2000). In a functional analysis, purified red seabream FSH failed to induce oocyte maturation in the red seabream, suggesting that LH plays an important role during early and late gametogenesis (Gen et al., 2003). During the growth of oocytes, follicle cells multiply and build a granulosa cell layer that surrounds the oocyte. Simultaneously, stroma cells form an outer flattened theca cell layer, less structurally organized than the inner granulosa layer. Both cell layers are important for the synthesis of steroidal mediators that are triggers for oocyte expansion and later maturation (Nagahama, 1994). Steroidogenesis is a complex process confined to specific steroidogenic cells involving the conversion of cholesterol to sexual steroids (Payne & Hales, 2004). Steroidogenesis is controlled by the action of GTHs that activate several signal transduction pathways. The most important is the cyclic-3’,5’-adenosine monophosphate (cAMP)/protein kinase A (PKA) pathway, but steroidogenesis is also stimulated by the protein kinase C (PKC) pathway, growth factors, and chloride and calcium ions (Stocco, Wang, Jo, & Manna, 2005). Cholesterol is the precursor not only for synthesis of sexual steroids in vertebrates, but also for mineralocorticoids (e.g., aldosterone) and glucocorticoids (e.g., cortisol) (H. Hsu, N. Hsu, Hu, & Chung, 2006). The steroidogenic acute regulatory (StAR) protein transports cholesterol from the cytoplasm to the inner mitochondrial membrane (Stocco, 2001), where the first steroidogenesisspecific enzyme, P450 side-chain-cleavage enzyme (P450scc), converts cholesterol to pregnenolone (Arukwe, 2008). Following this, several enzymatic pathways can be followed depending on the actions of 17a-hydroxylase/ 17,20b-lyase (P450C17), 20-hydroxysteroid dehydrogenase (20b-HSD), 17b-HSD, and other enzymes (for a graphical overview see Villeneuve et al., 2007). Synthesis of T takes place in theca cells, which release the hormone to granulosa cells, which then convert T into E2 (Nagahama, 1994), which is responsible for stimulating Vtg synthesis in the liver and leads to oocyte growth. There is growing evidence that androgens play a growth-promoting role in oocytes, at least in some species of teleosts (Kortner, Rocha, & Arukwe, 2009a; 2009b). Both 11-KT and T induce growth in previtellogenic oocytes

Chapter | 4


of Atlantic cod (Gadus morhua), which suggests a role of androgens in early follicular and oocyte growth. Gene expression patterns of oocytes differ in response to exposure to T and 11-KT and reveal novel androgen-responsive genes (Kortner, Rocha, Silva, Castro, & Arukwe, 2008). In previtellogenic eel oocytes, 11-KT induces an increase in oocyte diameter and stimulation of development, an effect not observed for E2 (Lokman, George, Divers, Algie, & Young, 2007). In the same study, a stimulating effect on oocyte diameter is also found for insulin-like growth factorI (IGF-I). Another important role for IGF-I is suggested during early vitellogenesis in the sterlet (Acipenser ruthenus), and IGF-I and the IGF-I receptor mRNA increased in cohorts of follicles from different maturation stages (Wuertz, Gessner, Kirschbaum, & Kloas, 2007; Wuertz et al., 2007b). There is evidence that other parameters promote early previtellogenic oocyte growth, suggesting an interaction between the growth axis and the reproductive axis. Larger Pacific salmon during a given reproductive developmental period possessed advanced stages of oocytes compared with smaller fish (Campbell et al., 2006). The same study showed a positive relationship between pituitary and plasma FSH, plasma E2, and ovarian StAR protein transcript level during the cortical alveoli accumulation period, whereas during the following lipid droplet accumulation period pituitary and plasma FSH, plasma E2, ovarian StAR protein, IGF-I and IGF-II transcript levels, and plasma IGFI were increased. The clear relationships among these endocrine factors suggest that mechanisms regulating body growth may influence the growth of previtellogenic oocytes. Some or all of those factors may be involved in the recruitment of oocytes into vitellogenesis, analogous to the situation of follicular selection in mammals (Fortune, Rivera, Evans, & Turzillo, 2001; Chandrashekar, Zaczek, & Bartke, 2004). The marked enlargement in size during oocyte secondary growth is mainly attributed to vitellogenesis. Uptake of Vtg in the oocytes and its conversion into yolk proteins is a particularly important process since the deposition of yolk in the oocytes builds the basic nourishment of the developing zygote after fertilization. Further, proteolysis of yolk proteins is involved in regulating the specific buoyancy of mature eggs, an important feature particularly for pelagic eggs of marine teleosts. Proteolysis leads to free amino acids that, together with ions, are involved in hydration of the eggs and adjusting their buoyancy (Carnevali et al., 2006). 17b-estradiol is synthesized by cooperation of the theca and granulosa cell layers surrounding the oocytes, and subsequently E2 is secreted into the blood. 17b-estradiol induces the synthesis of Vtg in the liver by entering through diffusion, binding and activating the ER. The dimeric occupied ER complex stimulates transcription of

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estrogenic genes including Vtg (Arukwe & Goksoyr, 2003). After protein synthesis and folding, Vtg is transported via the blood to the ovary, where it is taken up by oocytes via receptor-mediated endocytosis of Vtg receptor complexes at their membrane surface. Vitellogenin sequestered by endocytosis via surface receptors is cleaved into lipovitellins and phosvitins. Lysosomic enzymes play a key role in the cleavage of Vtg, especially cathepsin D (Carnevali, Carletta, Cambi, Vita, & Bromage, 1999; Kwon, Prat, Randall, & Tyler, 2001). Activation of cathepsin D by cathepsin B is probably necessary to achieve full activity, since the highest levels of both cathepsins were observed in early vitellogenic oocytes (Carnevali et al., 2006). After cleavage of Vtg, both products are stored as yolk protein. During vitellogenesis, several ER variants are present, as well as several Vtg genes (Mommsen & Koorsgaard, 2008). Besides the tentative assignment of the variants of ER to different physiological functions within fishes (Socorro, Power, Olsson, & Canario, 2000; Hawkins & Thomas, 2004; Filby & Tyler, 2005), the functions of gene variants in vitellogenesis have not been fully explored. However, it has been demonstrated that different forms of Vtg are processed, incorporated into fish oocytes, and differentially exploited by the embryo (Reith, Munholland, Kelly, Finn, & Fyhn, 2001; Sawaguchi, Ohkubo, Koya, & Matsubara, 2005). Several Vtg genes originated as a result of teleost genome duplication (Finn & Kristoffersen, 2007), and neofunctionalization has been suggested for these Vtg genes. The free amino acid pool within the oocyte cytoplasm that affects osmolarity and allows oocytes to float is derived by differential use of Vtg units and results mostly from proteolysis of the VtgAa paralog (Finn & Kristoffersen, 2007). Further, neutral lipids are produced by the activity of lipoprotein lipase (LPL), especially in midand late vitellogenesis, when lipid globules are incorporated into oocytes (Ibanez, Peinado-Onsurbe, Sanchez, & Prat, 2003). High amounts of neutral lipids serve as a caloric reserve for embryonic development. Lipoprotein lipase was localized in the follicle cells, and its activity is likely involved in building up oocyte lipid reserves (Ibanez, Peinado-Onsurbe, Sanchez, Cerda-Reverter, & Prat, 2008). The brain provides neuroendocrine regulation for the onset of puberty and oogenesis. Puberty is defined as the developmental process leading to the achievement of full reproductive capacity of a sexually immature fish. In female fish this means the period from the beginning of vitellogenesis until ovulation of mature eggs. The mechanism leading to the onset of puberty is not yet clear but it is suggested that genetic factors, metabolic signals, and environmental stimuli are involved. In mammals, hypothalamic GnRH activity is low between birth and puberty, and reactivation of GnRH release is regarded as a key event for the onset of puberty (Terasawa & Fernandez, 2001).

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The GnRH system integrates molecular and endocrine signals and stimulates GTHs that regulate ovarian development (Nocillado & Elizur, 2008) and function. In teleosts, two or three variants of GnRH are present, derived from distinct genes (Lethimonier, Madigou, Munoz-Cueto, Lareyre, & Kah, 2004) and, depending on the number of variants, either GnRH1 or GnRH3 is the hypophysiotropic isoform (Sherwood & Wu, 2005). Puberty can proceed if the stimulatory signals regulating GnRH activity prevail over the inhibitory signals. Several neurotransmitters have a modulatory activity on the GnRH system in fishes: g-aminobutyric acid (GABA), neuropeptide Y (NPY), norepinephrine, and serotonin are stimulatory, whereas dopamine (DA) acts as an inhibitor (Senthilkumaran, Okuzawa, Gen, & Kagawa, 2001). Effects of these neurotransmitters on the preoptic-anterior hypothalamus are observed during immaturity, recrudescence, and spawning in red seabream (Senthilkumaran et al., 2001). At the beginning of vitellogenesis, GnRH variants, GPR54 (a G-protein-coupled receptor for kisspeptin) and DA receptor showed highest expression in the brain and, during active vitellogenesis, DA receptor levels decreased in the pituitary, while GnRH1 and GPR54 increased in the ovary of grey mullets (Mugil cephalus) (Nocillado, Levavi-Sivan, Carrick, & Elizur, 2007). These results suggest that dopaminergic inhibition and positive regulatory signals of GPR54 and GnRH1 could be involved in early regulation of vitellogenesis. In some fishes, DA is inhibitory to reproductive processes, including puberty, as seen in various species kept in captivity (Aizen, Meiri, Tzchori, LevaviSivan, & Rosenfeld, 2005). Treatment of mullet females with a DA antagonist accelerates oocyte development and growth, raises E2 levels, and induces vitellogenesis. These results are in line with a dopaminergic inhibition of GnRH secretion and indicate that, by overcoming this inhibition, E2 levels increase due to stimulatory action of GTHs. A striking example of the inhibitory role of DA was shown during the onset of puberty in the European eel (Vidal et al., 2004). Combined treatment with T, a GnRH agonist, and a DA receptor antagonist stimulated vitellogenesis, indicating that the removal of dopaminergic inhibition in prepubertal European eels allows GnRH to stimulate LH release and cause ovarian development (Vidal et al., 2004). The onset of puberty in the European eel provides a good model for studying which environmental cues and endogenous factors are necessary to remove DA inhibition (Dufour et al., 2005). Recent evidence shows that DA could negatively modulate cyp19b expression in the female grey mullet brain (Nocillado et al., 2007). Teleost fishes are known to have very high cyp19b expression levels in the brain, which are further upregulated by E2 via an estrogen response element (ERE) (Callard, Tchoudakova, Kishida, & Wood, 2001). The balance between the stimulatory role of E2 and the


inhibitory role of DA on GnRH activity could be one of the triggers for early oocyte growth, taking into account that plasma E2 levels increase markedly during the course of this developmental period. Accordingly, in the early stage of puberty, cyp19b expression is minimal in the grey mullet (Nocillado et al., 2007). On the other hand, in Nile tilapia, DA receptor is upregulated by E2 (Levavi-Sivan, Aizen, & Avitan, 2005). An ERE on the promoter sequence of DA receptor was not verified, but alternate sites for regulatory transcription factors such as Sp-1 or NFkB were found (Nocillado, Levavi-Sivan, Avitan, Carrick, & Elizur, 2005). Detection of a significant role for GPR54 receptor and its ligands has rapidly revolutionized our view of gonadal development, in that all players have been identified. The impact of the GPR54 receptor on reproductive function was initially suggested after observing hypogonadotropic hypogonadism in humans and mice with an inactivating mutation in this receptor (De Roux et al., 2003; Seminara et al., 2003). This contention was confirmed by a growing body of literature that revealed a role for the KiSS-1 system (the GPR54 receptor and its ligands, kisspeptins encoded by the KiSS-1 gene) in control of reproductive processes in males and females. The KiSS-1 system is involved in stimulating the GnRHeGTH axis, in the control of puberty, and it has also been proposed as the key intermediary element in regulating positive and negative feedback effects of sex steroids on GTH secretion (Roa & TenaSempere, 2007). Our knowledge of the KiSS-1 system was obtained in mammals and its role in nonmammalian species mainly remains to be explored. However, some evidence indicates that the KiSS-1/GPR54 system may be involved in the control of reproduction in fishes. Fluctuations during puberty of GPR54 mRNA in the brain and ovary of grey mullet were observed, with highest expression of GPR54 and GnRH in the brain at the onset of puberty (Nocillado et al., 2007). Expression of GPR54 has been described in GnRH neurons from immature and mature male tilapia (Parhar, Ogawa, & Sakuma, 2004). During immaturity, a high percentage of GPR54 transcripts were detected in GnRH3 neurons, whereas in mature animals they were found in GnRH1, GnRH2, and GnRH3 neurons. In the fathead minnow (Pimephales promelas), KiSS-1 receptor (¼ GPR54 receptor) expression was highest in the brain at the onset of puberty, in females (at the cortical alveolus stage) and in males (when spermatogonia type B appear), and was considerably lower in the following stages (Filby, Van Aerle, Duitman, & Tyler, 2008). In the same study, it was shown that mammalian kisspeptin-10 not only stimulated expression of the KiSS-1 receptor, but also of GnRH3. This stimulatory role of kisspeptin on GnRH is known in mammals, and is attributed to an increase of GnRH release at the activation of puberty (Han et al., 2005; Castellano et al., 2006b).

Chapter | 4


5. OOCYTE MATURATION AND OVULATION A major event that follows vitellogenesis and oocyte growth is the acquisition of oocyte maturation competence (OMC), a process regulated by actions of GTHs. Oocytes that are arrested in prophase I of meiosis, during their growth phase, reinitiate meiosis, visible as GVBD. This maturational phase is dependent on LH, and is characterized by a preovulatory surge of LH that triggers the advancement of the oocytes to metaphase II. Synthesis of the maturation-inducing hormone (MIH)dor maturationinducing steroiddis required for oocyte meiotic resumption, and the ability of the follicle-enclosed oocyte to respond to the maturational signal of MIH is defined as OMC. Actions of LH on oocytes are mediated or modulated by a local paracrine network of peptide factors (Figure 4.3). In several teleosts, including salmonids (Nagahama, 1997) and tilapines (Rocha & Reis-Henriques, 1996; Rocha & Reis-Henriques, 1998), 17a,20b-dihydroxy-4pregnen-3-one (17a,20b-DP) has been identified as MIH. This compound is indeed regarded as the main MIH in fish, and the cooperation of granulosa and theca cells is responsible for synthesizing this steroid. Under the

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influence of GTHs, theca cells produce 17a-hydroxyprogesterone, which is in turn converted by granulosa cells to 17a,20b-DP (Nagahama, 1994). In contrast, 17a, 20b, 21-trihydroxy-4-pregnen-3-one (17,20b,21P) was identified as the MIH, for example in the Atlantic croaker (Micropogonias undulatus) (Trant, Thomas, & Shackleton, 1986) and in the sea bass (D. labrax) (Rocha & Reis-Henriques, 1999; 2000), both marine perciforms. The main enzyme responsible for producing MIH (17a,20b-DP) is 20-b-hydroxysteroid-dehydrogenase (20b-HSD). A steroidogenic shift is needed to proceed from vitellogenic growth towards oocyte maturation. Therefore, steroid synthesis has to be modulated from mainly E2 synthesis during vitellogenic growth towards the MIH production needed for oocyte maturation. The steroidogenic shift is possibly controlled by transcriptional factor binding sites, observed in the specific promoter regions of the genes (Senthilkumaran, Yoshikuni, & Nagahama, 2004). Both genes need the action of GTHs to increase their activity: ovarian aromatase via the Ad4BP/SF-1 motif, and 20b-HSD via the CREB motif. Their temporal expression patterns actually confirm their specific roles during the steroidogenic shift, with highest cyp19a1 and Ad4BP/SF-1 transcript levels

FIGURE 4.3 Overview of the physiological regulation of oocyte maturation in fishes. Luteinizing hormone (LH) is a key player during oocyte maturation that increases the activity of the maturation-inducing hormone receptor (MIH-R), via a still-unknown mechanism. Subsequently, MIH can bind to MIH-R and induce the maturation of oocytes. The LH signal is mediated by a local paracrine network that additionally triggers maturation. Insulin-like growth factor-I (IGF-I) increases the number of gap junctions and can independently induce the maturation of oocytes via the IGF-I receptor. The preovulatory surge of LH is triggered by positive feedback on GnRH in the brain. It is not known whether the KiSS-1 system is involved in this process in fishes as it is in mammals. 17a-OHP, 17a-hydroxyprogesterone; cAMP, cyclic-3’,5’-adenosine monophosphate; DA, dopamine; E2, 17b-estradiol; EGF, epidermal growth factor; GJ, gap junctions; GnRH, gonadotropin-releasing hormone; PKA, protein kinase A; PKC, protein kinase C; T, testosterone. See color plate section.

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during vitellogenesis and distinct expression of 20b-HSD around the time of spawning (Senthilkumaran et al., 2004). A two-stage model for oocyte maturation under control of GTHs has been proposed for fishes, and is consistent with at least ten teleost species and one amphibian species (Patino et al., 2001). The first stage comprises the acquisition of the ability of follicle cells to produce MIH as well as the ability of oocytes to respond to MIH. The second stage is the actual period of MIH production and the resulting resumption of meiosis. It is now well established that LH induces the production of MIH and sensitizes the oocytes to respond to MIH. Earlier studies showed that follicles previously primed with human chorionic gonadotropin (hCG) had an increased sensitivity to the maturational action of MIH (Jalabert, Breton, Brzuska, Fostier, & Wieniawski, 1977; Kime et al., 1987). In seabream, increased sensitivity to MIH was only obtained with LH treatment, and not with FSH (Kagawa, Tanaka, Okuzawa, & Kobayashi, 1998). This suggests that oocyte maturation is under the control of LH, and not part of the follicular growth actions of FSH. The induction of OMC by hCG in ovarian follicles, as monitored by GVBD onset, could be strongly reduced by exposure to inhibitors of transcription and translation, but not by an inhibitor of steroid synthesis. Hence, the acquisition of OMC is not mediated by steroids that could be enhanced by action of LH, but requires instead de novo transcription and translation (Patino & Thomas, 1990b). The action of LH may be mediated by second messengers. In similar experiments, adenylate cyclase activators and cAMP analogs stimulated OMC, whereas it was blocked by PKA inhibitors (Chang, Patino, Thomas, & Yoshizaki, 1999). Thus, the PKA-dependent signal transduction pathway is associated with the first stage of follicle maturation. Candidates for the suspected de novo transcription as mediators of the LH signal to achieve OMC could be 20b-HSD (the responsible enzyme for MIH production) or connexins and MIH receptor (important for gap junctions and reception of MIH signals, respectively). The MIH receptors are localized on the surface of oocytes and their activity is increased by LH during the first stage of maturation (germinal vesicle migration), followed by a decrease at ovulation. This evidence was observed in fishes and in amphibians (e.g., Patino & Thomas, 1990a; Liu & Patino, 1993) and indicates that MIH receptor activity is dependent on LH. Therefore, the LH-induced sensitivity of oocytes to MIH via the increase in the activity of MIH receptor(s) could play a significant role during acquisition of OMC. Another important aspect of follicle maturation is the participation of heterologous gap junctions as mediators of LH signaling (Yamamoto & Yoshizaki, 2008). Receptors of LH are only found on granulosa cells and not on oocytes (Nagahama, 1994); therefore, LH signaling has to


be transferred to the oocytes. Gap junctions are intercellular channels that are built by hexamers of connexin protein subunits. Molecules with low molecular weight up to 1 kDA, including second messengers and ions, can pass freely between cells through these gap junctions (Yamamoto & Yoshizaki, 2008). Heterologous gap junctions are formed between different cell types (e.g., granulosa celleoocyte), whereas homologous gap junctions are formed between the same cell types (e.g., granulosa cellegranulosa cell). In at least some fishes the number of heterologous as well as homologous gap junctions increases during the period of OMC (Patino & Kagawa, 1999). In experiments that induced GVBD in oocytes by stimulation with hCG, the stimulatory effect could be blocked by a gap junction-specific inhibitor in a concentration-dependent manner (Yamamoto, Yoshizaki, Takeuchi, Soyano, & Patino, 2008). This study demonstrated the important role of gap junctions during acquisition of OMC and further experiments confirmed the involvement of PKA. The mechanism could be LH binding to LH-R on granulosa cells, which in turn transfer cAMP via the gap junctions to the oocyte (Yamamoto et al., 2008). In oocytes, cAMP activates PKA, which, via a presently unknown mechanism, leads to the activation of the MIH-R at the surface of oocytes. Thereby, oocytes gain the ability to be responsive to the maturational signal of MIH and consequently advance to OMC (Yamamoto & Yoshizaki, 2008). The induction of OMC is particularly important in fishes with asynchronous modes of spawning in that, at a given time, only some selected clusters of oocytes are triggered to reach the maturation stage. In synchronous spawners the acquisition of OMC is not as precise, and an overlap is sometimes observed between the FSH-dependent growth phase and the LH-dependent maturation phase. Other factors are involved in modulation of the maturation of follicles. Insulin-like growth factor-I is a potent player that induces OMC, but by a different pathway than via stimulation of MIH production. Oocyte maturation in vitro can be induced by hCG and by IGF-I independently, and a combined exposure of either compound with actinomycin D (an inhibitor of transcription) only inhibits acquisition of OMC in the case of hCG. Results have shown that IGF-I is involved in stimulation of follicular maturation (Kagawa, Kobayashi, Hasegawa, & Aida, 1994). Insulin-like growth factor-I or hCG alone increase the number of heterologous and homologous gap junctions in incompetent follicles (i.e., before acquisition of OMC) of the red seabream. This result is physiologically relevant since the number of gap junctions increases during the normal maturation of oocytes in teleost fishes (Patino & Kagawa, 1999). Further, in Xenopus, oocytes possess IGF-I receptors, and meiotic resumption is a direct effect of insulin or IGF-I on IGF-I receptors at the surface of oocytes

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(Hainaut, Giorgetti, Kowalski, Ballotti, & Van, 1991). The major source of IGF-I production is the liver, but there is also local production in the gonads, specifically in granulosa and theca cells (Schmid, Naf, Kloas, & Reinecke, 1999; Berishvili, D’Cotta, Baroiller, Segner, & Reinecke, 2006). These results together suggest an important stimulatory role of IGF-I for induction of oocyte maturation, in addition to the already mentioned positive influence on previtellogenic oocytes. In contrast, the neurotransmitter serotonin inhibits MIH-induced oocyte maturation and could be a physiological modulator of OMC (Cerda, Subhedar, Reich, Wallace, & Selman, 1998). An inhibitory effect of serotonin on the maturational signal of MIH was effective in late vitellogenic and full-grown follicles but decreased in follicles in an early stage of maturation. Recently, the concept that a local paracrine network in the fish ovary mediates or modulates the actions of GTHs has emerged. Peptide growth factors may play especially important roles, and it has been suggested that not only do follicle cells support and nurture oocytes, but active oocyte signaling may influence follicle development (Ge, 2005), as in mammals (Matzuk, Burns, Viveiros, & Eppig, 2002). The activin, inhibin, and follistatin system is one part of the local paracrine network in vertebrates. Actions of activin are often modulated by follistatin and antagonized by inhibin. Activin acts through activin type I and type II receptors, and in case of the former the activin binding stimulates intracellular proteins called Smads (Mathews, 1994; Pangas & Woodruff, 2000). Two major roles have been described for activin, one during the maturation of follicles and the other during OMC development. Activin could be one of the mediating factors involved in GTH signaling from the follicle cells to the oocytes. Two variants of activin exist (activin A and B), and they are thought to play different roles during follicle development. Activin B increases the rate of maturation of postvitellogenic follicles in zebrafish, and this effect can be blocked by follistatin (Pang & Ge, 1999). During acquisition of OMC, the stimulatory effect of LH on oocytes, to enable them able to respond to MIH, can be mimicked by activin A and B with similar potency (Pang & Ge, 2002). Production of activin A and follistatin can be stimulated by hCG (or goldfish pituitary extract) via the classical cAMPePKA pathway, whereas activin B is suppressed by GTHs via a cAMPdependent but PKA-independent pathway (Wang & Ge, 2003a; 2003b). Temporal expression patterns suggest that activin A and follistatin are involved in promoting follicle and ovarian growth, whereas activin B could play an active role during ovulation (Wang & Ge, 2004b). Both activins are predominantly expressed in follicle cells, but their specific receptors are present on the oocytes, suggesting that oocytes are direct targets for activin actions. Follistatin is expressed more in oocytes than in follicle cells and probably its main role is to modulate the action of activin.

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Additionally, it suggests that the oocyte actively participates in follicle development (Wang & Ge, 2003c). Another paracrine factor in the teleost ovary is epidermal growth factor (EGF). Epidermal growth factor is involved in promoting oocyte maturation (Pati, Balshaw, Grinwich, Hollenberg, & Habibi, 1996) and in suppressing atresia in preovulatory follicles (Janz & Van der Kraak, 1997). Epidermal growth factor receptor expression increases during follicle development, and, interestingly, EGF is expressed in both follicle cells and oocytes, but the EGF receptor is solely present in follicle cells (Wang & Ge, 2004a). This suggests that EGF is a paracrine signal from the oocytes that targets the follicle cells and may regulate functions of the somatic follicle layers. Epidermal growth factor interacts with the aforementioned activinefollistatin system by upregulating both activins A and B, and by downregulating follistatin. As a consequence, an intrafollicular feedback loop has been proposed that involves a reciprocal regulatory mechanism (Ge, 2005). Follicle cells signal via activin on the activin receptor-expressing oocytes (stimulating intracellular Smad signaling proteins), while oocytes signal via EGF on EGF receptor-expressing follicle cells. Other local paracrine factors also may have profound roles in regulating oocyte development, but their significance is not yet fully explored. Pituitary adenylate cyclaseactivating polypeptide (PACAP) induces cAMP, and three PACAP receptor variants are present in zebrafish (Miyata et al., 1989; Wang, Wong, & Ge, 2003; Fradinger, Tello, Rivier, & Sherwood, 2005). Transforming growth factorb (TGF-b) suppresses both GTH and 17a, 20b-DP-induced oocyte maturation in zebrafish (Kohli et al., 2003), and growth differentiation factor-9 (GDF-9) is regarded in mammals as an oocyte-specific factor that controls folliculogenesis by influencing transcriptional activity in follicle cells (Matzuk et al., 2002). However, such a role is unknown in fishes. Generally, oocyte development is primarily controlled by GTHs, but there exists a complex, local regulatory network in the ovary to mediate or modulate their actions. Indeed, extensive communications have been described among the distinct compartments of the follicle complex: the oocyte and its surrounding somatic follicle cells. It is well known that some follicles mature completely, while others undergo follicular atresia. The degree of follicular atresia is dependent on the species and can comprise between 75e99.9% of follicles. In mammals, atretic apoptosis is initiated by the granulosa cells, whereas in fishes theca cells appear to be responsible (Wood & Van der Kraak, 2001; 2002). Lysosomal enzymes likely play a key role during follicular atresia; e.g., the proteases cathepsin B and L. In repeated-spawning teleosts, postovulatory follicles regress by apoptosis during ovarian recovery after spawning (Carnevali et al., 2006).

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Mature eggs are released to the gonoducts by a process called ovulation, in which oocytes separate from their surrounding follicle cells. Ovulation in mammals is a cyclical process and triggered by the preovulatory surge of GTHs. Neuroendocrine regulation of ovulation involves a rise in E2 by the dominant follicles and a subsequent increase in GnRH secretion. The KiSS-1 system is involved in generating the preovulatory surge of LH in mammals and therefore has an important role during ovulation (Roa & Tena-Sempere, 2007). During this period, LH secretion is activated by kisspeptins. Interestingly, in the hypothalamus two populations of KiSS-1-expressing neurons were found, one located at the arcuate nucleus (ARC) and the other at the anteroventral periventricular nucleus (AVPV). At the ARC, neuronal KiSS-1 mRNA is decreased by E2 and mediates the negative feedback effect of ovarian steroids on GTH secretion. At the AVPV, neuronal KiSS-1 mRNA is positively regulated by E2 and therefore could play a major role in the generation of the preovulatory surge of LH (Smith, Popa, Clifton, Hoffman, & Steiner, 2006). Recent data demonstrate that KiSS-1 and GPR54 are also expressed in the rat ovary in a cyclic-dependent manner (Castellano et al., 2006a). In immature animals, levels of KiSS-1 mRNA were very low, but they could be enhanced by GTH priming. The functional relevance of these findings remains to be defined, but the ability of the LH surge to induce KiSS-1 mRNA expression in the ovary suggests a potential role in controlling ovulation (Roa & TenaSempere, 2007). Interestingly, in ovaries of fishes the expression of GPR54 has been reported in the grey mullet (Nocillado et al., 2007) and in the fathead minnow (Filby et al., 2008), thus suggesting that a similar evolutionarily conserved mechanism could be present in fishes. Further research is needed to establish a role of the KiSS-1/GPR54 system in the regulation of reproduction in teleosts. Finally, inhibition by DA is involved in the regulation of ovulation in fishes. Mullets kept in captivity undergo vitellogenesis, but do not reach the spawning stage. Injections of these mullets with GnRH and a DA antagonist significantly increase the spawning rate above that induced by GnRH alone (Aizen et al., 2005). The participation of DA in the regulation of ovulation is a general feature of teleosts, as demonstrated in salmonids and cyprinids. Dopamine exerts direct inhibitory effects on gonadotropes, in contrast to the stimulatory actions of GnRH (Dufour et al., 2005). After ovulation, there is a wide diversity of breeding strategies in fishes regarding the fate of fertilized eggs. Many fish species just release mature oocytes without taking any further care. After fertilization, some embryos develop within their egg case at the bottom of the water body whereas others are pelagic until the larvae hatch, migrating to a specific depth according to their buoyancy. Some fish species hide their eggs; for example, salmonids


lay them in nests buried in sediments. Another strategy to increase the survival of offspring is parental care that is normally observed as egg guarding or as fanning of the nest to supply oxygen (Lissaker & Kvarnemo, 2006). Parental care is particularly advanced in mouth-breeding species that provide protection for their offspring in their own mouth, as known from African cichlids (e.g., Pseudocrenilabrus multicolour) (Corrie, Chapman, & Reardon, 2008).

6. FINAL CONSIDERATIONS Reproductive development and its physiological regulation in female fishes, as discussed and reviewed in this chapter, have received great scientific interest during recent decades. Basic principles underlying the regulation of ovarian development have emerged and seem to be evolutionarily conserved among species and within vertebrates. Future research on ovarian development and function in teleosts should focus on the early stages of oocyte development, especially on the maintenance and fate of primordial germ cells and on their local recruitmentdwe know very little about it. The same applies to the factors and mechanisms governing oocyte atresia. Further, integrating the molecular bases of cellular differentiation and functions, as well as putting together a coherent model of signaling networks during all stages of oocyte development, are promising yet challenging lines of research. Teleosts are especially suited as model animals due to their wide variety of strategies to differentiate their sex, and to develop and mature their oocytes (e.g., gonochoristic or hermaphroditic species, group-synchronous or asynchronous oocyte maturation forms). Comparative approaches among different teleosts should offer a great potential to get new insights into the factors that are responsible for gonadal differentiation, oocyte development, and maturation. The identification of genes, endocrine, and paracrine factors involved in the regulation of different strategies of oocyte development and maturation, and the analyses of their specific functions, may be of particular value for both biomedical and aquaculture research. Females of some fish species kept in captivity or used in aquaculture production often experience reproductive failures or delayed maturation of germ cells (Mylonas & Zohar, 2001). Yet there are crucial gaps in our knowledge of mechanisms leading to the formation of a viable egg. A greater understanding of reproductive physiology in those species will support the development of strategies to enable their controlled reproduction in captivity, an important prerequisite for their use in aquaculture. This latter aspect is particularly important since wild populations of marine and freshwater fishes are currently decreasing, and expansion of aquaculture is regarded as one future alternative, exploring

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the utilization of new species and improving production conditions.

ABBREVIATIONS 11-KT 17a-OHP 17a,20b-DP 17,20b,21P 20b-HSD 20b-HSD AMH ARC AVPV cAMP DA E2 EGF ER ERE FSH GABA GDF-9 GSI GTH GV GVBD hCG HPG IGF-I LH LPL MIH NPY OMC P450C17 P450scc PACAP PGC PGO PKA PKC StAR T TGFb Vtg

11-ketotestosterone 17a-hydroxyprogesterone 17a,20b-dihydroxy-4-pregnen-3-one 17a, 20b, 21-trihydroxy-4-pregnen-3-one 20-b-hydroxysteroid-dehydrogenase 20-hydroxysteroid dehydrogenase Anti-Mu¨llerian hormone Arcuate nucleus Anteroventral periventricular nucleus Cyclic-3’,5’-adenosine monophosphate Dopamine 17b-estradiol Epidermal growth factor Estrogen receptor Estrogen response element Follicle-stimulating hormone g-aminobutyric acid Growth differentiation factor-9 Gonadosomatic index Gonadotropin Germinal vesicle Germinal vesicle breakdown Human chorionic gonadotropin Hypothalamicepituitaryegonadal Insulin-like growth factor-I Luteinizing hormone Lipoprotein lipase Maturation-inhibiting hormone Neuropeptide Y Oocyte maturation competence 17a-hydroxylase/17,20b-lyase P450 side-chain-cleavage enzyme Pituitary adenylate cyclase-activating polypeptide Primordial germ cell Primary growth oocyte Protein kinase A Protein kinase C Steroidogenic acute regulatory Testosterone Transforming growth factor-b Vitellogenin

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Chapter 5

Thyroid Hormones and Reproduction in Fishes Jason C. Raine University of Saskatchewan, Saskatoon, SK, Canada

SUMMARY Thyroid hormones (THs) appear to be involved in various aspects of growth and development in fishes, but their potential role in reproduction is not clear. A large number of published studies have attempted to show a relationship between THs and the reproductive cycle in fishes, but differences in species and their reproductive strategies, and methods used to analyze changes in TH regulation and signaling, have likely contributed to conflicting results. There is evidence to suggest that changes in thyroid function do take place during the reproductive cycle, but whether these changes reflect a direct role in reproductive events or merely the corresponding changes in energy utilization, decreased growth, and other factors accompanying the onset of the reproductive cycle is not apparent. This chapter reviews the current understanding of the regulation and signaling pathways of THs and their potential involvement in fish reproduction.

1. INTRODUCTION Thyroid hormones (THs), the iodinated thyronines triiodothyronine (T3) and thyroxine (T4), are thought to be involved in a large number of biological processes in fishes, including metabolism, growth, development, and reproduction. However, very few definitive roles have actually been attributed to THs. In many cases, the literature is rife with experimental evidence both substantiating and challenging various proposed functions of THs. In mammals, maintenance of metabolic homeostasis could be considered the major function of THs, and likely the best known. Thyroid hormones are also involved in a number of physiological and developmental processes, and the involvement of T3 in many aspects of mammalian metabolism and temperature regulation is well established. The role of THs in thermogenesis likely evolved in concert with endothermy, and it is evident that thermogenesis is closely tied to metabolic regulation. This suggests that THs were involved in metabolic regulation prior to the evolution of


endothermy, and that the role of THs changed with the development of endothermy. Thus, there is a possibility that metabolic processes in poikilotherms involve THs, even though metabolic rates in these animals are primarily driven by ambient temperature. Although a number of studies have described effects of THs on metabolism in fishes, the evidence is generally conflicting, and the doses of T4 or T3 administered in many of these studies are in the pharmacological range (see Leatherland, 1994, for review). Nevertheless, the possibility exists that T3 plays multiple permissive roles in metabolic regulation in fishes, as it does in mammals. Thyroid hormones appear to play a major role in vertebrate development. In particular, a large body of evidence suggests that THs are strongly associated with neural development and maturation. In mammals, T3 is essential for neuronal proliferation, migration, synaptogenesis, and myelination during brain development (Howdeshell, 2002; Horn & Heuer, 2010). Developmental abnormalities such as mental retardation, deaf-mutism, and motor disorders can result from hypothyroidism in the developing human fetus, but these problems can be partially reversed by TH replacement after birth (Hulbert, 2000). Further, in cases where the mother or developing fetus are hypothyroid, there is neurological impairment of the fetus (congenital cretinism), and the degree of impairment is correlated with the level of hormone insufficiency (LaFranchi, 1999). In fishes, neurological effects of THs also occur. Hindbrain formation in zebrafish (Danio rerio) embryos is impaired when the TH receptor (TR) isoform, TRa1, is overexpressed (Essner, Johnson, & Hackett, 1999). Moreover, THs induce programmed cell death (apoptosis) of ultraviolet-sensitive (UVS) cone cells and corresponding loss of sensitivity to ultraviolet (UV) light over most of the neural retina of rainbow trout (Oncorhynchus mykiss) during development (Browman & Hawryshyn, 1992; 1994; Kunz, Wildenburg, Goodrich, & Callaghan, 1994; Deutschlander, Greaves, Haimberger, & Hawryshyn, 2001; Allison et al., 2003; 83

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Hawryshyn, Martens, Allison, & Anholt, 2003; Allison, Dann, Veldhoen, & Hawryshyn, 2006; Kunz, 2006; Veldhoen et al., 2006; Raine & Hawryshyn, 2009; Raine, Coffin & Hawryshyn, 2010). One well-studied aspect of TH functions involves the events surrounding organ restructuring in poikilothermic vertebrates, either during embryonic development or during metamorphosis. Thyroid hormones control the initiation of metamorphosis of tadpoles to adult amphibians, and are directly associated with many tissue-specific changes that take place during this metamorphic process (see Buchholz, Paul, Fu, & Shi, 2006; Brown & Cai, 2007, for review). Similarly, flatfishes of the order Pleuronectiformes undergo a metamorphic transformation during development that is directly regulated by THs. These flatfishes transform from a bilaterally symmetrical, pelagic larval stage into a benthic, asymmetrical juvenile. Major behavioral, physiological, and morphological changes take place in these fish during metamorphosis and this process can be stimulated with the addition of exogenous T4, or inhibited with the addition of goitrogens, drugs that inhibit production of endogenous THs (Miwa & Inui, 1987; Power et al., 2001). Thyroid hormones also may play a role in other developmental stages of fishes that are not considered metamorphic, but do involve substantial changes in morphology, behavior, and physiology (Hoar, 1988; Lam, 1994; Leatherland, 1994; Power et al., 2001; Campinho, Silva, Sweeney, & Power, 2007). These include the transition of embryonic/ larval fish stages that feed exclusively from yolk reserves containing THs of maternal origin, to the free-feeding juvenile stage as well as smoltification, the transformation of salmonid fishes (trout and salmon) during migration from shallow, freshwater streams to the ocean. In all vertebrates, a role for THs in reproduction has been investigated, yet the function of THs, if any, in this process is elusive. In mammals and birds, THs have been linked to changes in day length and the initiation of seasonal reproduction, although the mechanism by which THs act is unknown (Nakao, Ono, & Yoshimura, 2008). In fishes, a role for THs in reproduction has been sought for more than 50 years, and there is still no clear answer. Many fish studies have examined the reproductive cycle with a dominant regulatory function for THs in mind, and evidence that THs could be involved in the process of gonadal maturation in some species has emerged. It may be that, in searching for a major controlling role of THs in reproduction, more specific and understated roles of THs have been overlooked. Further, our increasing knowledge of TH regulation and action suggests that classical means of analyzing TH action and involvement in reproduction may need to be reassessed. This chapter reviews the current understanding of TH physiology and the relationship between THs and reproduction in fishes.


2. THYROID HORMONE DELIVERY 2.1. Regulation of Circulating Thyroid Hormone (TH) Levels The concentration of THs in the blood is generally maintained within a relatively narrow range, especially in mammals where THs are tied to metabolic rate and thermogenesis. The regulatory pathway responsible for maintaining stable levels of circulating THs is referred to as the hypothalamicepituitaryethyroid (HPT) axis (Figure 5.1). The model for TH regulation is similar for all vertebrates, but in mammals it begins with the release of thyrotropin-releasing hormone (TRH) from the hypothalamus. Thyrotropinreleasing hormone is a tripeptide that is highly conserved across all vertebrate groups, although its functions at least with respect to TH homeostasis seem to be dissimilar (Zoeller, Tan, & Tyl, 2007). In mammals, TRH stimulates the release of thyrotropin (TSH) from thyrotropes in the anterior pituitary gland (Zoeller, Tan, & Tyl, 2007). Thyrotropinreleasing hormone is transported directly from the hypothalamus to the adenohypophysis region of the pituitary gland through the hypophysial portal blood system, which is found in mammals (Zoeller et al., 2007). Teleost fish lack this portal blood system and instead the adenohypophysis is directly innervated by hypothalamic neurosecretory fibers (Peter, Yu, Marchant, & Rosenblum, 1990; Leatherland, 1994). In mammals, TRH controls blood levels of TSH through binding to a TRH receptor on a thyrotrope, which results in activation of the phosphatidyl inositol-protein kinase C pathway, and initiates the synthesis and release of TSH (Rondeel et al., 1988; Bogazzi et al., 1997). Also in

Hypothalamus ?

Pituitary

T4 T3

Blood Vessel

TSH +

T4 T3

T3 T4

Liver MD Kidney Gill

Thyroid Tissue

FIGURE 5.1 Simplified diagram demonstrating the regulation of plasma thyroid hormones (THs) in fishes through the hypothalamicepituitarye thyroid (HPT) axis. Release of thyrotropin (TSH) from the anterior pituitary stimulates thyroxine (T4) and lesser amounts of triiodothyronine (T3) to be synthesized and released from the thyroid tissue. Most of the T3 circulating in the bloodstream comes from deiodination of T4 by monodeiodination enzymes (MD) in peripheral tissues. High plasma TH levels decrease TSH release through negative feedback. The regulation of TSH release may involve an unidentified hypothalamic factor.

Chapter | 5

Thyroid Hormones and Reproduction in Fishes

mammals, somatostatin (SS) inhibits TSH secretion in a similar manner and acts as a negative control of TH levels in the blood (Vtiger, 1987). Although several studies have attempted to show the effects of TRH on either TSH secretion or T4 and T3 levels in fishes, the results regarding the possible involvement of SS and TRH in thyroidal control have been ambiguous (see Leatherland, 1994, for review). Recently, isolated pituitary cells from bighead carp (Austichthys nobilis) have been shown to increase TSH mRNA levels after exposure to TRH (Chatterjee, Hsieh, & Yu, 2001). Additionally, cell-surface TRH receptors have been identified in a fish pituitary and they are structurally similar to those of mammals (Harder et al., 2001). In anurans, corticotropinreleasing factor (CRF), not TRH, appears to stimulate TSH release, and there is some evidence that CRF could play a similar role in fishes (Larsen, Swanson, Dickey, Rivier, & Dickhoff, 1998; Okada et al., 2007). However, there is also evidence that the release of TSH from the thyrotropes of the pituitary gland of some teleost fishes may be under inhibitory hypothalamic control, and this has been demonstrated by hypothalamic lesioning and pituitary transplantation studies (see Leatherland, 1994, for review). However, the potential hypothalamic factors responsible for this inhibitory action have still not been fully identified in fishes. In teleost fishes, as in mammals, TSH appears to be the main factor regulating TH release from the thyroid follicles (Figure 5.1). Thyrotropin is one of three glycoprotein hormones in the pituitary gland that share a common alpha subunit (glycoprotein hormone subunit a (GPHa)) together with a beta subunit that determines the hormone’s biological activity; i.e., the TSHb subunit for TSH (Zoeller, Tan, & Tyl, 2007). Sequencing of the beta subunit of the TSH gene in several fish species has demonstrated a highly conserved DNA sequence (Ito, Koide, Takamatsu, Kawauchi, & Shiba, 1992; Pradet-Balade, Schmitz, Salmon, Dufour, & Que´rat, 1997; 1998; Martin, Wallner, Youngson, & Smith, 1999; Yoshiura, Sohn, Munakata, Kobayashi, & Aida, 1999; Chatterjee, Hsieh, & Yu, 2001; Wang, Zhou, Yao, Li, & Gui, 2004; Lema, Dickey, Schultz, & Swanson, 2009). In several of these species, TSHb expression was detected in the pituitary and a number of other tissues including the liver, kidney, testis, and ovary (Wang et al., 2004; Lema et al., 2009). Extra-pituitary expression of TSHb was unexpected and current research is aimed at understanding the functional significance of this finding. In the last several years, TSH receptor (TSH-R) cDNAs have been cloned from tissues of the striped bass (Morone saxatilis) (Kumar et al., 2000), amago salmon (Oncorhynchus rhodurus) (Oba, Hirai, Yoshiura, Kobayashi, & Nagahama, 2000; 2001), sunrise sculpin (Pseudoblennius cottoides) (Kumar & Trant, 2001), African catfish (Clarias gariepinus) (Vischer & Bogerd, 2003), European sea bass (Dicentrarchus labrax) (Rocha et al., 2007), and channel catfish (Ictalurus punctatus) (Goto-Kazeto, Kazeto, & Trant,

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2009). Thyrotropin receptors are 30 kDa dimeric proteins transmembrane G-protein-coupled receptors belonging to the glycoprotein hormone receptor family, which includes the luteinizing hormone (LH) receptor (LH-R) and folliclestimulating hormone (FSH) receptor (FSH-R) (Vassart, Pardo, & Costagliola, 2004). Thyrotropin receptors are expressed solely in thyrocytes of amago salmon, whereas TSH-R expression has been found to be abundant not only in thyroid tissue but also in gonads of the striped bass, European sea bass, sunrise sculpin, African catfish, and channel catfish (Kumar et al., 2000; Kumar & Trant, 2001; Vischer & Bogerd, 2003; Rocha et al., 2007; Goto-Kazeto et al., 2009). Thyrotropin receptors transcripts have also been detected in various other tissues of these fishes, including skeletal muscle, heart, brain, liver, and kidney (Kumar et al., 2000; Vischer & Bogerd, 2003; Rocha et al., 2007; Goto-Kazeto et al., 2009). The presence of TSH-R in thyroid tissue provides further support for the regulatory role of TSH in thyroid function, whereas the presence of TSHb-subunit and TSH-R in extra-thyroidal tissues suggests that TSH possibly has some as yet unknown functions. Administration of TSH increases plasma T4 concentrations and increases thyroid tissue mass and activity in several species of teleost fishes (see Smith & Grau, 1986; Leatherland, 1994; for review see Raine, Takemura, & Leatherland, 2001). Thyrotropin increases iodide uptake by thyrocytes and iodination of tyrosyl residues (Zoeller et al., 2007). Thyrotropin also increases releases of T4 and T3 from thyrocytes in both rainbow trout and medaka (Oryzias latipes) (Raine et al., 2001) by upregulating the expression of genes involved in TH synthesis and release (Vassart & Dumont, 1992; De Felice, Postiglione, & Di Lauro, 2004). In the majority of in-vivo studies carried out in fishes, TSH does not directly alter plasma T3 levels, even though plasma T4 levels are elevated (Leatherland, 1994). Similarly, a corresponding decrease in conversion of T4 to T3 commonly occurs when plasma T3 levels are elevated following the administration of exogenous T3 (Eales et al., 1990; Eales & Finnson, 1991). This provides strong evidence for the independent regulation of plasma T4 and T3 levels (Leatherland, 1994; Leiner & MacKenzie, 2001). In all vertebrates, relatively high levels of circulating T4 and T3 initiate negative feedback mechanisms that result in decreased T4 and T3 release from the thyroid tissue and reduced circulating T4 and T3 levels (Figure 5.1). In this way, organismal plasma T4 and T3 levels are maintained within a reasonably narrow range. In mammals, in addition to stimulating TSH release, TRH is thought to modulate the sensitivity of the pituitary to negative feedback by THs (Greer, Sato, Wang, Greer, & McAdams, 1993). Both T4 and T3 negatively regulate the synthesis and release of TSH in the mammalian pituitary and indirectly affect TSH synthesis through changes in TRH synthesis (Bogazzi et al.,

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1997). In both mammals and teleosts, THs decrease TSH synthesis and release by binding to nuclear TRs in the thyrotropes. Triiodothyronine is thought to bind to thyrotrope TRs whereas T4 acts indirectly through intra-pituitary and intra-hypothalamic T4 to T3 conversion by monodeiodinating enzymes (Bogazzi et al., 1997). Nevertheless, both T4 and T3 directly decrease transcription of the TSHbsubunit gene in both the mammalian and teleost pituitary (Bogazzi et al., 1997; Pradet-Balade, Schmitz, Salmon, Dufour, & Que´rat, 1997; 1998). Recently, human pituitary thyrotropes have been shown to contain TSH receptor transcripts, suggesting the existence of a negative feedback loop enabling downregulation of TSH transcription in the presence of high TSH levels (Prummel et al., 2000; Theodoropoulou et al., 2000; Zoeller et al., 2007). Three main forms of iodinated thyronines are found in the blood of fish: T4, T3, and the inert metabolite of T4, reverse-triiodothyronine (rT3) (Figure 5.2). Thyroxine is the main hormone released from the thyroid tissue of teleost fishes, while T3 is released in lesser amounts (Chan & Eales, 1975; Eales, 1979; Grau et al., 1986; Byamungu, Mol, & Kuhn, 1992; Raine & Leatherland, 1999; 2000; Raine et al., 2001). Traditionally, T4 was considered a prohormone for T3, which was thought to be the biologically active form of TH, due to the much higher affinity of nuclear TRs for T3. However, the recent discovery of a cell surface TR specific for T4 in mammals suggests that this paradigm may need to be re-evaluated (Bergh et al., 2005). Deiodinating enzymes (deiodinases) located primarily in the liver as well as in TH target tissues are important regulators of vertebrate TH concentrations in the blood and provide either an increased peripheral supply of T3 or the clearance of both T4 and T3 from the circulation (Eales & Brown, 1993; Leatherland, 1994; Ko¨hrle, 1999; Bianco, Salvatore, Gereben, Berry, &

T4

I

O

I

HO I

OH NH2

O I

Outer ring deiodination

T3 HO I

Inner ring deiodination

I

OH NH2

O I

rT3

I

O

O

HO I

OH NH2

O I

FIGURE 5.2 The chemical structure of the main thyroid hormones in vertebrates showing the position of iodine atoms on the phenolic (tyrosine) rings. Removal of an iodine atom from the outer ring of thyroxine (T4) by deiodination enzymes creates triiodothyronine (T3), while removal of an iodine atom from the inner ring results in reverse T3 (rT3).

Larsen, 2002; Hernandez & St. Germain, 2003; Brown et al., 2005; Nunez, Celi, Ng, & Forrest, 2008) (Figure 5.1). Local tissue deiodination is considered an important mechanism for supplying T3 in the nervous system, and T3 levels in a target cell can be increased or decreased by deiodination (Darras, Mol, Van der Geyten, & Kuhn, 1998; Nunez, Celi, Ng, & Forrest, 2008). Enzymatic monodeiodination of T4 to T3 involves the removal of one iodine atom from the outer ring of the molecule (Leatherland, 1994; Mol et al., 1998). Reverse T3 is generated by the removal of one iodine atom from the inner ring of the T4 molecule (Figure 5.2). The liver, gill, and kidney are responsible for the majority of the deiodination (Figure 5.1), although many other tissues have deiodinase activity (Byamungu et al., 1992; MacLatchy & Eales, 1992; Morin, Hara, & Eales, 1993; Mol et al., 1998). Three types of deiodinase enzyme have been identified thus far in mammals, birds, amphibians, and fishes, namely type I, type II, and type III. In mammals, type I deiodinase is thought to be found in all tissues, but it has the highest activity in the liver, the kidney, thyroid tissue, and the central nervous system (CNS) (Hulbert, 2000). Type I deiodinase can perform both inner- and outer-ring deiodination, is inhibited by propylthiouracil (PTU), and is responsible for generating the majority of the circulating T3 via conversion of T4 into T3. It also generates rT3 from T4 (Yen, 2001). Type II deiodinase performs outer-ring deiodination only, and is not inhibited by PTU. It has been identified in the CNS, brown adipose tissue, anterior pituitary gland, and placenta (Hulbert, 2000). Type II deiodinase functions predominately to convert T4 into T3 to supply the local intracellular needs for T3 (Yen, 2001). Type III deiodinase performs only inner-ring deiodination (converting T4 into rT3 and T3 into the inactive diiodothyronine, T2). It is found in the CNS, placenta, and skin and also is not inhibited by PTU (Hulbert, 2000; Yen, 2001). Less is known about the monodeiodinases of fishes relative to mammals, but deiodinase types I, II, and III have been found in a number of fish species (see, e.g., Mol et al., 1998), and cDNAs for all three deiodinase enzyme types have been isolated from killifish, (Fundulus heteroclitus) and tilapia (Oreochromis niloticus) (Valverde, Croteau, Lafleur, Orozco, & St. Germain, 1997; Sanders et al., 1999; Orozco, Villalobos, & Valverde, 2002), which suggests that the deiodinases have been highly conserved among vertebrate classes. However, the type I deiodinase in teleosts appears to possess a number of functional characteristics, including an insensitivity to PTU and altered responsiveness to increased T3 levels, that suggest that this enzyme is not as similar to its mammalian counterpart as once thought (Van der Geyten, Byamungu, Reyns, Kuhn, & Darras, 2005). Several studies have shown that teleost type I deiodinase activity is not affected and gene expression is downregulated during hyperthyroidism, in contrast to the

Chapter | 5

response of this enzyme observed in mammals (Berry, Kates, & Larsen, 1990; Mol et al., 1998; Plohman et al. 2002; Garcıá, Lopez-Bojorquez, Nunez, Valverde, & Orozco, 2004). In addition, hepatic type I deiodinase activity and expression is upregulated in hypothyroid tilapia, again disparate to the response of this enzyme seen in mammals (Berry et al., 1990; Van der Geyton, 2001).

2.2. Thyroid Hormone (TH) Transporters Thyroid hormones exist in two forms in the blood: free and bound to transport proteins (Figure 5.3). Less than 1% of the T4 and T3 present in the blood of teleosts is estimated to be present in the free form, and thus readily accessible to target cells (Eales & Shostak, 1985; Weirich, Schwartz, & Oppenheimer, 1987). The remainder of the hormone is

Blood Vessel T3

87


T4

T3 Alb

T4

T4

TTR

Alb

T4

T4

T4

T4R? MD T4

T3

2.3. Thyroid Hormone (TH) Clearance

T3 TR

reversibly bound to transport proteins, of which albumin was thought to be the primary form in teleosts until transthyretin (TTR) cDNA was identified in the gilthead sea bream (Sparus auratus) (Santos & Power, 1999). The T4binding globulin (TBG) present in mammals has not been detected in fishes. Both TTR and albumin in mammals have a higher affinity for T4 than T3, whereas in fish, birds, and amphibians TTR has a higher affinity for T3 than for T4 (Richardson, 2009). In addition to the liver, TTR transcripts have been detected in the heart, skeletal muscle, kidney, testis, gills, and pituitary of adult gilthead sea bream, and in the liver of several salmonids during smoltification (see Richardson, 2009). Recent studies in mammals suggest that there are specific transporters responsible for transfer of THs into target cells. These transmembrane transporters appear to be Naþ-independent organic anion transport systems, which transport THs across the membranes of target cells in the CNS, liver, kidney, and retina (Heuer & Visser, 2009). They may also transport conjugated and unconjugated steroid hormones from the bloodstream into hepatocytes (Abe, Suzuki, Unno, Tokui, & Ito, 2002). Although there is no direct evidence for similar TH transmembrane transporters in fishes, indirect evidence suggests that T4 and T3 transporters are present in teleosts. Thyroxine and T3 entry into red blood cells occurs by simple diffusion, as well as by pHand temperature-sensitive saturable transport mechanisms (McLeese & Eales, 1996a; 1996b). There also is evidence for separate T4 and T3 transporters. Transport of T3 into red blood cells is Naþ-independent and relatively rapid, whereas T4 uptake is slower, very specific, and appears to depend on binding to red blood cell protein (McLeese & Eales, 1996b).

RXR

TRE

gene Nucleus

Target Cell FIGURE 5.3 Thyroid hormone action at the cellular level in teleost fishes. Thyroid hormones are present in the blood bound to the plasma binding proteins, albumin (Alb) and transthyretin (TTR), or in free form. Bound thyroxine (T4) and triiodothyronine (T3) are inaccessible to target tissues and provide a circulating store of hormone, while the free form is available to enter target cells via a specific cellular transporter. Alternatively, T4 may bind to a specific receptor on the cell membrane as in mammals. Triiodothyronine may be transported directly into the cell, but it is postulated that mainly T4 in the cell provides the substrate for T3 generation through deiodination enzymes (MD). Triiodothyronine enters the nucleus where it binds to its receptor already associated with a thyroid response element (TRE) on the target gene. Thyroid hormone receptor (TR) usually binds as a heterodimer with a retinoid X receptor (RXR).The TR alone represses transcription, while T3 binding to the receptor initiates transcriptional machinery to generate messenger RNA. MAPK, mitogenactivated protein kinase.

In mammals, THs are conjugated in the liver through sulfation/sulfonation by sulfotransferases, or glucuronidation by UDP-glucuronosyl transferase prior to elimination through the bile (Hood & Klaassen, 2000a; 2000b). These enzymes alter the solubility of THs and prevent deiodination (Zoeller et al., 2007). In teleosts, a similar clearance mechanism appears to exist. All of the principle thyronines (T4, T3, and rT3) undergo glucuronidation and sulfation in rainbow trout in vivo, and by hepatocytes in vitro (Finnson & Eales, 1996; Finnson, McLeese, & Eales, 1999). Glucuronides of T3 occur in the blood of male tilapia (DiStefano, Ron, Nguyen, Weber, & Grau, 1998). Deiodination also serves to clear T4 and T3 from the blood and tissues of teleosts (DiStefano et al., 1998).

2.4. Thyroid Hormone (TH) Receptors Thyroid hormones produce their effects primarily after binding to two classes of TR: a nuclear receptor with

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a higher affinity for T3 than for T4 and a plasma membrane receptor that has an affinity primarily for T4. Thyroid nuclear receptors have a genomic mechanism of action whereas the plasma membrane receptor works mainly through kinase cascades.

2.4.1. Nuclear triiodothyronine (T3) receptors In mammals, THs readily enter target cells and exert most of their effects at the genomic level via their interaction with the TR in the nucleus of target cells (Figure 5.3). Intracellular type I or II deiodinases can convert T4 into T3, which then binds to the TR. The occupied TR may bind to the thyroid response element (TRE) on DNA as a homodimer but binds most strongly as a heterodimer with the retinoid X receptor (RXR) (Hulbert, 2000; Yen et al., 2006; Mengeling et al., 2008; Koury et al., 2009). In the absence of ligand, TRs are thought to bind to DNA as either a homodimer or a heterodimer with RXR, and stimulate the binding of a corepressor complex that actively represses basal transcription of the target gene (Hulbert 2000; Harvey & Williams, 2002). Two key corepressor proteins that frequently make up the corepressor complex with TRs are the silencing mediator of retinoid and TH receptor (SMRT) and the nuclear receptor corepressor (NCoR), found in both mammalian and amphibian systems (Tomita, Buchholz, & Shi, 2004; Choi et al., 2008). This model is the generally accepted method of T3dependent gene activation. In vertebrates, two highly conserved forms of TR, TRa and TRb, have been identified, each encoded by a separate gene (Harvey & Williams, 2002). Two additional TR transcripts have been identified for each of the TRa and TRb proteins in mammals: TRa-1 and TRa-2, and TRb-1 and TRb-2. TRa-1 and TRa-2 arise from alternate splicing of the initial RNA transcript, while TRb-1 and TRb-2 are generated through the use of one of the two promoters located on the TRb gene (Yen, 2001). TRa-2 is the only isoform that does not bind T3. A number of other TRa and TRb subtypes have also been identified in mammals and fishes, although their functional significance is still being investigated (Zoeller et al., 2007; Nelson & Habibi, 2009). TRa-1 and TRb-1 are expressed in most tissues, but their relative abundance varies (Harvey & Williams, 2002). In fishes, TRa and TRb mRNA have been identified in a number of species, including the Japanese flounder (Paralichthys olivaceus) (Yamano, Araki, Sekikawa, & Inui, 1994; Yamano & Inui, 1995), halibut (Hippoglossus hippoglossus) (Llewellyn et al., 1999), gilthead sea bream (Nowell, Power, Canario, Llewellyn, & Sweeny, 2001; Power et al., 2001), rainbow trout (Jones, Rogers, Kille, & Sweeney, 2002; Raine, Cameron, Vijayan, Lamarre, & Leatherland, 2004), zebrafish (J. Essner, Breuer, R. Essner, Fahrenkrug, Hackett, 1997; Liu, Lo, & Chan, 2000),


Atlantic salmon (Salmo salar) and tilapia (Marchand et al., 2001), conger eel (Conger myriaster) (Kawakami, Tanda, Adachi, & Yamauchi, 2003a; 2003b), turbot (Scophtalmus maximus) (Marchand, Duffraisse, Triqueneaux, Safi, & Laudet, 2004), goldfish (Carassius auratus) (Nelson & Habibi, 2006), and Pacific bluefin tuna (Thunnus orientalis) (Kawakami, Nozaki, Seoka, Kumai, & Ohta, 2008). TRa and TRb mRNA has been detected in most tissues of the Japanese flounder, zebrafish, and other species, although the isoforms are generally differentially expressed in different species (Yamano & Miwa, 1998; Liu et al., 2000; Nelson & Habibi, 2009). Nuclear TRs also are present in oocytes and early embryos of zebrafish and rainbow trout, fertilized eggs of Japanese flounder, and blastula-stage embryos of gilthead sea bream (Essner et al., 1997; Yamano & Miwa, 1998; Liu et al., 2000; Power et al., 2001; Raine et al., 2004; Li, Raine, & Leatherland, 2007). Receptor binding and signaling by T3 in fishes is highly conserved relative to that found in mammals and other vertebrates (e.g., see Nunez et al., 2008; Nelson & Habibi, 2009). Thyroid hormones upregulate their own nuclear TRs in mammals, amphibians, and fishes, as shown in a number of studies using adult, juvenile, and embryonic developmental stages. Fathead minnows (Pimephales promelas) fed a T3enhanced diet increased both TRa and TRb transcripts in the liver and brain (Lema et al., 2009). Similarly, exogenous T4 exposure through immersion or body cavity implant consistently upregulated retinal TRa and TRb gene expression in rainbow trout parr during the TH-dependent developmental process of UVS cone degeneration (Raine & Hawryshyn, 2009; Raine et al., 2010). In amphibians and several fish species, including the Japanese flounder, Senegalese sole (Solea senegalensis), and conger eel, that undergo TH-dependent metamorphosis, increases in whole body TH levels precede metamorphic climax and correspond to increased accumulation of TR transcripts (Yamano & Miwa, 1998; Kawakami et al., 2003a; 2003b; Tata, 2006; Manchado, Infante, Rebordinos, & Canavate, 2009). Both TRa and TRb gene expression were upregulated in response to exogenous T4 and T3 treatment in conger eel hepatocytes cultured in vitro (Kawakami et al., 2006), whereas T4 treatment upregulated TRb expression in larval Senegalese sole and treatment with a goitrogen (thiourea) decreased TRb transcripts (Manchado et al., 2009). The findings of these studies suggest that TH upregulation of TRs may be a necessary phenomenon that exists to sensitize target tissues/cells to T3 in response to temporal tissue-specific requirements.

2.4.2. Plasma membrane thyroxine (T4) receptor It has been recognized for some time that not all actions initiated by THs are mediated through nuclear TRs, and the existence of a nongenomic TH signaling pathway was

Chapter | 5


Blood vessel Aereolar connective

tissue

Lumen

Thyroid follicle

Thyrocyte showing T4 immunostaining

Periphery of lumen and colloid exhibiting T4 immunostaining

Blood vessel

suspected. In fishes, TH-induced rapid stimulation of succinate dehydrogenase in hepatic mitochondria of carp (Cyprinus carpio) and the potential enhancement of gonadotropin-induced estradiol (E2) secretion in trout oocytes was thought to take place too quickly for T3 to bind to its nuclear receptor and initiate transcription (Cyr & Eales, 1989; Peter & Oommen, 1989). Many more examples of nongenomic actions of THs can be found in mammals, including effects on mitochondria, glucose uptake, and actin polymerization, the latter of which promotes neural outgrowth and may be a key factor in TH effects on brain development (Shih et al., 2004; Farwell & Leonard., 2005; Moeller, Cao, Dumitrescu, Seo, & Refetoff, 2006; Mousa, O’Connor, F. Davis, & P. Davis, 2006; Zoeller et al., 2007; Casas et al., 2008; Leonard, 2008; Lombardi et al., 2009). A plasma membrane receptor for T4, the integrin vb3 receptor, was recently identified in mammals and it appears to be the source of these nongenomic effects of THs (see P. Davis, Leonard, & F. Davis, 2008, for review). This membrane receptor has a higher affinity for T4 than T3, and binding of T4 to the receptor activates a mitogen-activated protein kinase (MAPK) signal transduction pathway in angiogenesis studies (Davis et al., 2009). Although a cell membrane-associated T4 receptor has yet to be identified in any fish, it may be that this is also the source of a number of nongenomic effects in this vertebrate group too. It is clear that a re-evaluation of the classical model of TH action, where T4 is considered to be only a prohormone for T3, is in order.

3. THE THYROID TISSUE OF FISHES The thyroid follicle is considered to be the functional unit of the thyroid gland and is highly conserved in all vertebrate species. Thyroid follicles are made up of a single layer of epithelial cells (thyrocytes) creating an extracellular compartment or lumen containing colloid (Figure 5.4). The

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FIGURE 5.4 Thyroid tissue of juvenile rainbow trout immunostained for thyroxine (T4). This histological image shows the general appearance of the thyroid tissue in many species of teleost fish. Thyroid follicles are scattered throughout the areolar connective tissue of the lower jaw in close association with the blood supply. Thyroxine immunostaining demonstrates the presence of T4 in some of the thyroid epithelium cells (thyrocytes), the periphery of the colloid, and the lumen. This staining technique can be used to help assess the activity level of the thyroid tissue. An increase in the number of T4-immunostained thyrocytes, and increased intensity of staining in and around the colloid, indicate increased activity or storage. Processing of the tissue for histology can result in shrinkage or loss of the colloid from the lumen of the thyroid follicles, as seen in this image.

formation of an extracellular lumen is highly conserved among all vertebrates and is thought to be a required adaptation for TH synthesis, which involves a highly reactive oxidative step, and extracellular storage (Leatherland, 1994). It may be that, by sequestering this oxidative reaction in an extracellular compartment, the thyroid follicle provides protection for other tissues and cells. Mammals, birds, amphibians, and reptiles possess a discrete glandular thyroid surrounded by a connective tissue covering. Similarly, cartilaginous fishes also possess an encapsulated thyroid gland, but, in most species of bony fish, the thyroid tissue does not form a discrete gland but consists of isolated thyroid follicles scattered throughout the wellvascularized areolar connective tissue of the lower jaw, in the vicinity of the ventral aorta (Leatherland, 1994) (Figure 5.5). In teleost fishes, although seemingly active thyroid follicles are detected prior to hatch in a number of freshwater species, activity of the tissue is based solely upon histological criteria and corresponding increases in whole body TH levels have not been observed to corroborate this finding (Atlantic salmon (Hoar, 1939), fathead minnows (Wabuke-Bunoti & Firling, 1983) chinook salmon (Oncorhynchus tschawytscha), and coho salmon (Oncorhynchus kisutch) (Leatherland & Lin, 1975; Greenblatt, Brown, Lee, Dauder, & Bern, 1989)). Further, the pattern of active thyroid follicle appearance in marine species differs from that of freshwater fish, and active follicles are not found until after yolk absorption, again using only histological characteristics (Nacario, 1983; Brown et al., 1988). Immunohistochemical detection of T4 and T3 in the developing thyroid tissue of rainbow trout embryos demonstrated that, in these fish, synthesis of the THs begins prior to hatch and release of THs from the thyroid tissue does not occur until much later after hatch but before the onset of exogenous feeding (Raine & Leatherland, 1999; 2000). An example of T4-immunostained thyroid tissue from a juvenile rainbow trout can be seen in Figure 5.5.

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Thyroid Follicle

Lumen

T4

T4

T4

TG T4 T4

T4 TG

T4

T3

T3

morphology of thyroid follicles, the equivalent chemical structure of THs, and the similarities in regulation and signaling of TH suggest that synthesis and release of THs by thyroid follicles would be similar among all vertebrates, and studies in teleosts appear to corroborate this assumption (Eales & Brown, 1993; Leatherland, 1994; Cyr & Eales, 1996; Raine & Leatherland, 2000; Raine et al., 2001; Blanton & Specker, 2007; Nelson & Habibi, 2009). Details of the process of TH synthesis that applies to all vertebrates are provided in Figure 5.4.

Lysosome

T3 TPO

K+

Na+

Na+

TG 2Na+

ILumen

K+

I-

NIS

2Na+

I-

Thyrocyte Blood Vessel

FIGURE 5.5 Synthesis and release of thyroid hormone from thyroid tissue. Iodide (I-) is taken up by the thyroid epithelial cell (thyrocyte) by sodium iodide symporters (NIS), which rely on a sodium (Naþ) gradient established by sodium potassium ATPases (diamond). I- travels through the cell and likely enters the lumen of the follicle through a channel protein. The protein that makes up thyroglobulin (TG) is manufactured in the thyrocyte and is iodinated upon entering the thyroid follicle lumen by a thyroid-specific thyroid peroxidase (TPO) to form the insoluble thyroid hormone-associated TG that makes up the colloid. The stored TG is solubilized by proteases and taken up by the thyrocyte by endocytosis. Lysosomal proteases release the thyroid hormones from the TG molecule and mainly thyroxine (T4), with some triiodothyronine (T3), is released from the cell into the bloodstream.

Similarly to mammalian thyroid gland development, TSH does not appear to play a role in early thyroid tissue development of teleost fishes, although TSH RNA has been detected in unfertilized oocytes of channel catfish (I. punctatus) and could potentially be involved (Raine & Leatherland, 2000; Alt et al., 2006, Goto-Kazeto et al., 2009). Thyrotropin immunoreactive cells have been found prior to the detection of TH synthesis in the developing thyroid tissue of rainbow trout embryos, and may be involved in stimulating the onset of TH synthesis (Raine & Leatherland, 1999; 2000).

3.1. Thyroid Hormone (TH) Synthesis and Release Most of the information on TSH stimulation of TH synthesis and release can be found in studies of the mammalian thyroid gland. However, the highly conserved

4. THYROID HORMONE (TH) AND REPRODUCTION IN FISHES A possible role for THs in reproduction in fishes has been sought for several decades, but, despite the large body of literature published in this area, a clear function for THs in this process has not been revealed, and a role in reproduction is still unclear. There are many excellent reviews dealing with TH and reproduction in fishes that compile, synthesize, and discuss the main studies in this area prior to the 21st century, and these should be consulted for a comprehensive review of the earlier literature (see Eales, 1979; Leatherland, 1982; 1987; 1988; 1994; Cyr & Eales, 1996). The general consensus of these prior reviews exploring the role of THs in fish reproduction appears to be that, although there is evidence to suggest involvement of THs in reproduction, there is also a wealth of evidence suggesting that THs are not directly involved in the reproductive process, which is a pervasive theme when reviewing the literature and attempting to define a role for THs in fish. As has been discussed in other reviews, the basis of the conflicting results attained with much of this research may be attributed to the diverse methods used to investigate TH action, the variety of species examined, and the many different reproductive strategies that they employ. A great deal of the early studies on TH and reproduction involved the correlation of plasma T4 and T3 levels with the reproductive cycle of various species of fish. However, as discussed earlier in this review, it is now evident that TH levels are highly regulated both in the blood and at the tissue/cellular level. Moreover, it has become quite apparent that detection of a change in one measurement does not automatically infer a functional change in TH action. Changes in the regulation of THs can generally compensate and overcome disturbances to individual components of this regulatory system (Zoeller et al., 2007). Thus, there is a need to ensure that evaluation of TH physiology and action involves the measurement of multiple factors involved in TH regulation and action. Just as critical is the need to develop TH-specific biomarkers with clear functional endpoints that can be used to gauge the severity of perturbations to general TH physiology and

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regulation, and to determine whether this interference results in a real impact at the organismal level. A number of reviews have addressed this growing concern of how to accurately assess a change in thyroid function and this will not be directly addressed in this chapter (see Eales & Brown, 1993; Brown, Adams, Cyr, & Eales, 2004; Blanton & Specker, 2007). The more informative published studies that address the involvement of TH in fish reproduction fall into one of the following categories: (1) correlative studies examining changes in TH function during reproductive maturation; (2) identification of TH regulatory elements in gonadal tissue; (3) assessment of thyroid function following sex steroid treatment; or (4) manipulation of thyroid function to induce changes in reproductive maturation. These categories are employed in subsequent sections to aid in the review of TH in fish reproduction.

4.1. Correlative Studies Examining Changes in Thyroid Hormone (TH) Function During Reproductive Maturation A great many early studies have investigated the possibility that THs play a major role in controlling reproductive events. These investigations generally employed measurement of plasma TH levels and/or thyroid tissue histology to correlate changes in reproductive events that take place during the breeding cycle. A thorough review of this early body of work can be found in Cyr & Eales (1996). In general, there is a positive correlation between the activity level of the thyroid tissue and the reproductive stage of seasonally breeding, non-salmonid teleost fishes, despite variations in life history and habitat (Cyr & Eales, 1996). Thyroid tissue activity tends to increase during the early stages of gonadal development, and these levels are maintained or increased during reproduction, and decrease after spawning (Cyr & Eales, 1996). This pattern of thyroid activity also is seen in fish species with short breeding cycles where thyroidal status increases during the early stages of each cycle. Further, reproductive status and thyroid tissue activity appear to possess a clear temporal relationship in salmonids, based on thyroid histology and elevated plasma TH levels (Cyr & Eales, 1996). These fishes exhibit an increase in thyroid activity with the onset of gonadal development and a general decrease in plasma TH levels with the upstream spawning migration in both males and females. In non-anadromous species there is often an increase in plasma TH levels at spawning, especially in males, followed by an increase in thyroid activity with the continuation of growth (Cyr & Eales, 1996). Similarly, in another seasonal spawner, the burbot (Lota lota), elevated blood T3 has been observed prior to spawning and decreased blood T3 at spawning, although no

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change was seen in T4 (Mustonen, Nieminen, & Hyvrinen, 2002). An earlier study did find increases in blood T4 at spawning, especially in females (Hornsey, 1977). However, despite the large amount of correlative evidence to bolster this interpretation, changes in water temperature, day length, and other seasonal parameters greatly complicate the matter (Leatherland, 1994; Cyr & Eales, 1996). This can be seen in the brown bullhead (Ictalurus nebulosus) and channel catfish (I. punctatus), which do not exhibit any relationship between plasma TH and gonadal development, but in which plasma TH levels correlate with ambient temperature (Burke & Leatherland, 1983; MacKenzie, Thomas, & Farrar, 1989; Leatherland, 1994). In oviparous elasmobranchs, similar positive correlations of thyroid activity with reproductive stage can be found, with minimal thyroid function observed in immature females and maximal thyroid activity seen at the peak of egg development and vitellogenesis, although the elevation in male thyroid activity is less pronounced (Cyr & Eales, 1996). In the stellate sturgeon (Acipenser stellatus), high thyroid activity occurs along with gonadal maturation during the prespawning migration as well as at spawning and even shortly after spawning has concluded (Pickford & Atz, 1957; Cyr & Eales, 1996). The age at which sturgeon achieve reproductive maturity varies from approximately seven to twenty-five years, and they generally have four- to seven-year intervals between successive spawning cycles, depending on the species and sex of the fish (Sulak & Randall, 2002). Moreover, it has been suggested that female sturgeon do not commonly reach as great an age as previously thought, and thus may not spawn many times during their average lifespan (Sulak & Randall, 2002). The effects of environmental factors on TH and reproductive events have been examined in trout of similar age exposed to altered photoperiod and constant water temperature and ration, in an effort to separate these parameters from reproductive processes (Cyr, MacLatchy, & Eales, 1988). This experimental regimen generated plasma TH levels that were highest during early ovarian development, as found previously, and it was found that plasma THs decreased as E2 increased. This pattern was present irrespective of photoperiod. In a comparable study, Pavlidis, Dessypris, and Christofidis (1991) found low THs in female rainbow trout during spawning compared to nonspawning females. Triploid stinging catfish (Heteropneustes fossilis) possess an uneven number of chromosomes and exhibit irregular meiotic division resulting in reduced gonadal development, as compared with diploid individuals (Biswas et al., 2006). Thyroid hormone levels were similar in diploid and triploid fish until spawning when diploid females exhibited significantly reduced plasma total T4 and T3 levels relative to the triploid females. No differences were seen in plasma TH levels in diploid and triploid males

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with spawning (Biswas et al., 2006). During the spawning period, female diploid catfish exhibited an increase in thyroid gland activity assessed by histology, and a significantly higher oocyte content of T4 and T3 than triploid females. It was suggested that oocyte TH uptake is responsible for the decrease in plasma TH profiles during spawning in diploid females and thyroid gland activity is increased to compensate for the low plasma TH levels at this time (Biswas et al., 2006). Interestingly, a change in the visual system of anadromous salmonids accompanies sexual maturation and upstream spawning migration of these fishes. The UVS cones and sensitivity to UV wavelengths of light reappear in maturing salmonids, and are thought to be required for navigation to natal streams (Browman & Hawryshyn, 1994; Beaudet, Novales-Flamarique, & Hayashi, 1997; Hawryshyn, 2000; Novales-Flamarique, 2000; Hawryshyn et al., 2003). The regeneration of UVS cones accompanies the surge in plasma T4 seen during the reproductive cycle in these fishes, and T4 treatment precociously induces UVS cone reappearance and sensitivity to light in the UV spectra (Browman & Hawryshyn, 1994; Hawryshyn et al., 2003). Although this retinal change is not generally considered a reproductive event, it does consistently accompany the spawning migration in salmonids and is an event that during reproduction can be tied to THs. Although TH levels, and in some cases thyroid histology, often correlate quite well with the reproductive cycle in many fish species, particularly during early gonadal maturation, there are other physiological changes, such as changes in energy partitioning, that accompany this stage that should be taken into account (see Cyr & Eales, 1996). The general inhibition of thyroid function as the reproductive cycle progresses is considered to be reflective of a shift in energy partitioning, where somatic growth is decreased to allow available energy to be used for gonadal growth (Leatherland, 1994; Cyr & Eales, 1996). However, even if the primary role of THs during the reproductive cycle is metabolic, this function still contributes to successful reproduction and thus could still be considered a reproductive role.

4.2. Identification of Thyroid Hormone (TH) Regulatory Elements in Gonadal Tissue In the year 2000, the TSH-R gene was sequenced and its expression detected in the thyroid tissue of amago salmon (Oba et al., 2000). Interestingly, at the same time, TSH-R expression was also found in the gonadal tissue of striped bass (Kumar et al., 2000). A short time later, gonadal TSHR expression was confirmed in the testis of the African catfish, the ovaries of the channel catfish, and the gonads of European sea bass, with additional expression observed in


several other tissues (Vischer & Bogerd, 2003; GotoKazeto, Kazeto, & Trant, 2003; Rocha et al., 2007). In both the European sea bass (D. labrax) and the channel catfish, TSH-R expression levels increased with ovarian maturation, but differed in the onset of TSH-R downregulation either before or after spawning (Rocha et al., 2007; GotoKazeto et al., 2009). Similarly, TSH-R expression increased to a maximal level during early testes development in these fishes (Rocha et al., 2007; Goto-Kazeto et al., 2009). Surprisingly, TSHb subunit transcripts were also detected for gonadal tissues of the orange-spotted grouper (Epinephelus coioides) and the red-spotted grouper (Epinephelus akaara)din the ooplasm of maturing oocytes and in spermatogenetic cysts, but not in the ovarian follicular cells (Wang et al., 2004). The TSHb expression pattern in these groupers complements that of the TSH-R expression pattern found in the other fish species (Kumar et al., 2000; Goto-Kazeto et al., 2003; Vischer & Bogerd, 2003; Rocha et al., 2007). Extra-thyroidal activity of TSH has been suggested previously in mammalian systems. In various mammalian tissues, TSH-R has been found in ovaries, lymphocytes, thymus, pituitary, testes, kidneys, brain, adipose/fibroblast, heart, and bone (reviewed by Davies, Marians, & Latif, 2002; Abe et al., 2003; Aghajanova et al., 2009). Recently, granulosa cells have been shown to have TSH-R, and TSH has been shown to increase cAMP concentration in these cells in culture (Aghajanova et al., 2009). These studies suggest that TSH may be directly involved in some aspects of gonadal physiology, and that this hormone may do more than just stimulate TH release from the thyroid. Receptors for THs have been detected in all stages of rat and human testicular development (Jannini et al., 1990; Jannini, Ulisse, & D’Armiento, 1995; Cooke, Zhao, & Bunick, 1994; Wagner, Wajner, & Maia, 2008; 2009). Further, T3 regulates the maturation and growth of testes, and controls Sertoli cell and Leydig cell proliferation and differentiation during testicular development (Wagner et al., 2008). Human ovarian granulosa cells have TRs, and T4 treatment of granulosa cells in culture initiates MAPK activation within 10 minutes, suggesting activation of the membrane T4 receptor (Rae et al., 2007; Aghajanova et al., 2009). In fishes, TRs have been reported in Leydig cells of freshwater climbing perch (Anabas testudineus), gonadal tissues of the hermaphroditic black porgy (Acanthopagrus schlegeli), and the gonads of the gonochoristic goldfish and fathead minnow (Jana & Bhattacharya, 1993; Nelson & Habibi, 2006; Lema et al., 2009; K. An, M. An, Nelson, Habibi, & Choi, 2010). In addition to TRs, deiodination enzymes are present in the gonads of several vertebrate species. Boar seminal plasma contains type II deiodinase activity, and this enzyme in the testes is postulated to act as a local regulator of TH levels and to provide the predominant source of T3

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for the testes (Brzezinska-Slebodzinska, Slebodzinski, & Kowalska, 2000). The activity level of this deiodination enzyme increases in piglet testicular homogenates from one to four weeks after birth (Brzezinska-Slebodzinska et al., 2000). Moreover, deiodinase II and III expression is present in luteinized human granulosa cells and human ovarian thecal cells (Rae et al., 2007; Aghajanova et al., 2009). In fishes, type II deiodinase expression has been detected in the testes and ovaries of the rainbow trout, and high plasma T4 levels correspond to the maximal number of type II deiodinase transcripts detected in the testes (Sambroni et al., 2001). Additionally, the increase in type II deiodinase expression correlates with the onset of spermatogenesis and increases in plasma T4, which suggests local abundance of T3 (Sambroni et al., 2001). The detection of TRs and deiodinases in gonadal tissues is not surprising, as TRs are present in most vertebrate tissues. Their presence does suggest a function in transcriptional regulation of some genes, but, without further research, a specific function cannot be ascribed. However, the additional presence of TSH in reproductive and other tissues is unexpected and warrants further investigation.

4.3. Assessment of Thyroid Function Using Sex Steroid Treatment Treatment with sex hormones has generated little supporting evidence for the general trends found in earlier studies examining correlative changes in the HPT and the hypothalamicepituitaryegonadal (HPG) axes during reproductive maturation. There is a general trend in a number of fish species for E2 to suppress thyroid function, while THs decrease production of E2. Treatment with E2 decreases total and free plasma T3 and sometimes plasma T4 levels in a number of fish species, including freshwater European eels (Anguilla anguilla) (M. Olivereau & J. Oliverau, 1979; M. Olivereau, Leloup, De Luze, & J. Olivereau, 1981), mollies (Poecilia sp.) (Sage & J. Bromage, 1970), rainbow trout (Cyr et al., 1988; Flett & Leatherland, 1989a; 1989b), and masu salmon (Oncorhynchus masou) (Yamada, Hiriuchi, Gen, & Yamauchi, 1993). Further, E2 suppresses hepatic T3 production and plasma T3 levels in salmonids (Cyr et al., 1988; Flett & Leatherland, 1989a; 1989b; Mercure et al., 2001). Moreover, decreased thyroid epithelial cell heights, reflecting decreased thyroid activity, have been found in European eel and rainbow trout following E2 treatment, even when no changes in circulating T4 levels were observed, suggesting a decrease in T4 clearance (Olivereau et al., 1981; Leatherland, 1985). Similarly, E2 treatment decreases T4 clearance and secretion rates via decreased T4 to T3 conversion in rainbow trout and the Southern lamprey (Geotria australis) (Cyr et al., 1988; Flett & Leatherland, 1989b; Leatherland et al., 1990; Mercure

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et al., 2001) and decreases free plasma T3 and total T4 levels, perhaps due to some degree of reduction in T4 binding to plasma carrier proteins (Cyr & Eales, 1989; 1992) or possibly by a decrease in the availability of TR for T3 binding (Bres, Cyr, & Eales, 1990). This inhibitory role of estrogens was demonstrated recently, when treatment with a synthetic estrogen (ethinylestradiol) resulted in TRa and TRb downregulation in fathead minnow liver, although TRb was simultaneously upregulated in the ovary (Filby, Thorpe, Maack, & Tyler, 2007). Alternatively, E2 enhances thyroidal status in the climbing perch (A. testudineus), dwarf snakehead (Channa gachua), and striped dwarf catfish (Mystus vittatus) (Singh, 1968; 1969; Chakraborti, Rakshit, & Bhattacharya, 1983; Chakraborti & Bhattacharya, 1984; Bandyopadhyay, Banerjee, & Bhattacharya, 1991). Further, removal of the ovaries from climbing perch results in a depression of thyroid function and plasma T4 levels that can be reinstated with E2 treatment (Chakraborti et al., 1983; Chakraborti & Bhattacharya, 1984). In contrast, thyroid function in sockeye salmon (Oncorhynchus nerka) and rainbow trout does not respond to E2 treatment (Van Overbeeke & McBride, 1971; Milne & Leatherland, 1978), although perhaps the thyroid axis was already depressed due to fasting (Milne & Leatherland, 1980). Clearly, there is no uniform pattern, but these differences in the effects of E2 may be species-specific or may be due to variations in experimental design, such as dose used, nutritional status of subjects, developmental stage, water temperature, and the method used to assess thyroid function. Interestingly, androgen treatment generally produces results opposite to those of E2 treatment, and enhances thyroidal function in most teleosts examined (Cyr & Eales, 1996). Testosterone (T) or methyltestosterone (MT) have been found to increase plasma TH levels in dwarf snakeheads (Singh, 1968; 1969), rainbow trout (Hunt & Eales, 1979; Leatherland & Sonstegard, 1980; Leatherland, 1985; MacLatchy & Eales, 1988; Shelbourn, Clarke, McBride, Fagerlund, & Donaldson, 1992), coho salmon (O. kisutch) (Fagerlund, Hijman, McBride, Plotnikoff, & Dosanjh, 1980), masu salmon (Ikuta, Aida, Okumoto, & Hanyu, 1985; Yamada et al., 1993), and the guppy (Poecilia reticulata) (Schwerdtfeger, 1979). Exposure to MT also stimulates increased activity of thyroid tissue in medaka (Nishikawa, 1976). Testosterone treatment of both rainbow trout and Arctic char (Salvelinus alpines) resulted in increased T4 to T3 conversion, further supporting the positive correlation between T and thyroid function (Hunt & Eales, 1979; MacLatchy & Eales, 1988). However, no effect of androgen treatment on plasma TH levels or conversion of T4 to T3 has been reported in other studies (Milne & Leatherland, 1980; Leatherland, 1985; Yamada et al., 1993). Nevertheless, as with E2 treatment, differences in experimental design, dosing, water temperature, sex, and

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the nutritional status confound the interpretation of these experiments. The use of hypothalamic gonadotropin-releasing hormone (GnRH) and pituitary gonadotropin (GTH) to study the role of THs in reproduction has generated mixed outcomes. Some of the varied results may stem from the fact that only one GTH was identified in fish prior to 1989, and many studies employed injection of isolated fish pituitary GTH that could potentially have included one or both forms of GTH. The fish GTHs are designated GTH-I, considered homologous to mammalian FSH, and GTH-II, homologous to mammalian LH (Kawauchi & Sower, 2005). Thyroid epithelial cell height increases following treatment of gonadectomized sockeye salmon with salmon GTH (Donaldson & McBride, 1974) and both GnRH analogs and salmon GTH increase plasma T4 in female sea lampreys (Sower, Plisetskaya, & Gorbman, 1985). Both salmon GTH and ovine LH markedly increase plasma T4 levels in climbing perch, while ovine FSH has no effect (Chakraborti & Bhattacharya, 1984). Further, treatment with a GnRH analog increases plasma T3 levels but not T4 in immature rainbow trout (Plate et al., 2002). However, plasma TH levels are unchanged with salmon GnRH treatment of goldfish, and salmon GTH does not alter secretion of T4 in medaka (Cyr & Eales, 1996). Further, suppression of thyroid function resulting from removal of the ovaries in climbing perch is not reinstated with subsequent salmon GTH treatment (Chakraborti et al., 1983; Chakraborti & Bhattacharya, 1984).

4.4. Manipulation of Thyroid Function to Induce Changes in the Reproductive System A number of fish studies have approached elucidation of the relationship between reproduction and TH in another way. These studies employed altered thyroidal status to examine the reproductive system for correlative changes. Using the advantage of an encapsulated thyroid tissue in cartilaginous fishes, thyroidectomy of female dogfish sharks (Scyliorhinus canicula) has been shown to prevent development of vitellogenic ovarian follicles (Lewis & Dodd, 1974). Since most teleosts possess diffuse thyroid tissue that cannot be completely removed from the living fish, chemical goitrogens have been widely used to reduce the synthesis and release of THs from the thyroid tissue. To this end, the use of chemically induced hypothyroidism in a number of teleost fishes has generally resulted in impaired gonadal function (Leatherland, 1987; 1994; Cyr & Eales, 1996; Swapna & Senthilkumaran, 2007). There is evidence that some goitrogens, such as thiourea, are considered to have toxic effects on fishes, and this could interfere with the interpretation of many of the studies using these and other potentially toxic chemicals (Leatherland, 1994). However,


the relatively consistent trend of decreased GSI and gonadal development/maturation accompanying decreased thyroid function reported in many studies is difficult to ignore. Decreased TH production in male African catfish treated with thiourea prior to spawning results in decreased spermatozoa in the testicular lumen (Swapna et al., 2006). Further, lowered circulating THs in a number of Asian teleosts interfere with a variety of testicular processes during maturation of the testes, which resume with a return of the fish to a euthyroid state (Swapna & Senthilkumaran, 2007). Although there are other studies that have found no effect or a detrimental effect of THs on several testesrelated processes, reproductive stage and treatment dose are likely factors in the range of outcomes reported (Swapna & Senthilkumaran, 2007). Nevertheless, these studies do indicate that THs can affect maturation of the testes in teleost fishes and suggest that THs may be involved in milt production, and sperm viability and maintenance. Decreased thyroidal activity has been shown to accompany reduced ovarian function in rainbow trout and fathead minnows exposed to sublethal concentrations of thiocyanate (Ruby, Idler, & So, 1993; Lanno & Dixon, 1994). Similarly, inhibition of thyroid function using sodium ipodate injections decreases the gonadosomatic index (GSI) in rainbow trout, whereas treatment with T3-supplemented food increases GSI (Cyr & Eales, 1988a). Ovarian follicle development is impaired in African catfish following thiourea treatment, and T4 treatment increases the growth rate of ovarian follicles and the number of mature follicles present in the ovarian tissue (Supriya et al., 2005). In zebrafish, PTU administration increases the number of oocytes produced, but decreases the size of the mature oocytes (Van der Ven, Van den Brandhof, Vos, Power, & Wester, 2006). Thyroxine also stimulates ovarian maturation in immature goldfish, but has no effect on ovarian development in hypophysectomized adults, suggesting a pituitary factor is also involved (Hurlburt, 1977). In the guppy, T4 treatment decreases spawning interval and brood time (Lam & Loy, 1985). Further, T3 restores the responsiveness of oocytes to GTH in the female stellate sturgeon, with delayed sexual development resulting from decreased water temperature (Detlaf & Davydova, 1979). Treatment with T4 increases the action of salmon GTH and ovine LH in-vitro on the ovary of the climbing perch (Chakraborti & Bhattacharya, 1984). However, T4 only had this effect two hours prior to salmon GTH treatment. In rainbow trout, T3 and/or salmon GTH increase GSI after 21 days, and this effect is greater in trout treated with both T3 and GTH than with GTH treatment alone (Cyr & Eales, 1988a). Moreover, T3 consistently potentiates salmon GTH stimulation of gonadal steroid secretion in vitro (Hurlburt, 1977; Cyr & Eales, 1988b; Cyr et al., 1988; Soyano, Saito, Nagae, & Yamauchi, 1993). In rainbow trout, this effect has been found to be bimodal, with low T3 treatment

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stimulating E2 production and higher T3 concentrations suppressing E2 production (Cyr et al., 1988). In medaka, a similar effect has only been seen 32 hours prior to ovulation, even though a surge of T4 and T3 release took place at 12 hours preovulation (Soyano et al., 1993). Further, synthesis of GnRH is decreased with T3 treatment in tilapia (Parhar et al., 2000). Recently, fathead minnows fed a T3-elevated diet have exhibited significantly elevated TSHb transcripts in the testes, and TRb and GPHa transcripts have been shown to be significantly elevated in both the ovaries and the testes (Lema et al., 2009). Gonadal TRa expression in the fathead minnow gonads, however, did not change following T3 treatment. In mice, TRa1 and TRb appear to have opposite roles in the control of reproductive behavior (Forrest, Reh, & Ru¨sch, 2002). Estrogens stimulate female receptivity to mating, and THs interfere. Further, TRa1-deficient mice exhibit decreased mating behavior, whereas TRb-deficient mice show increased mating behavior (Forrest et al., 2002). Clearly, changes in thyroid function can affect reproductive events, but this, in and of itself, does not prove a direct role for TH in reproduction. It may be that effects on reproduction are indirect through permissive effects of TH on related components of the reproductive system, or as a result of an interrelationship between physiological changes incurred with altered TH availability and reproductive events.

5. CONCLUSIONS Although there is still no clearly defined role for THs in reproduction, there is evidence to suggest that THs are involved in the reproductive cycles of fishes. The strong correlation between THs and nutritional status suggests that energy partitioning is a key role for THs during the reproductive cycle, shifting the mobilization of energy reserves from growth to reproductive maturation. There is also evidence that THs play major roles in testicular and ovarian development. However, teleosts are an incredibly large and diverse group encompassing many very different life histories and reproductive strategies, which can make interpretation and comparison between species exceedingly difficult and perhaps unrealistic. Further, much of the research in this area has employed single-parameter assessment of thyroid function/physiology, namely plasma TH levels, and searched for an overarching regulatory role for TH during the reproductive cycle. The delivery of THs to their target cells is highly regulated through redundant and compensatory mechanisms to maintain TH signaling despite perturbations resulting in altered plasma TH (Zoeller et al., 2007). This new appreciation for preservation of TH action irrespective of plasma TH levels requires re-evaluation of previous dogma, and revision of experimental assessment of TH physiology and action in many

vertebrates, especially in fishes. This includes a more thorough investigative method for experiments required to interpret functional changes in TH physiology, where analysis of multiple players involved in TH signaling and regulation is essential. Further, it is highly probable that the control of reproductive events involves the interaction of multiple hormones and that TH is only one of many components in this process. The success of future research in this area would appear to benefit from an emphasis on the potential involvement of THs in the various tissue-/cellspecific changes that take place during the reproductive cycles of fishes, rather than the overall regulation of this event. Evaluation of tissue/cellular regulation of THs, standardization of methods of analysis, and an understanding of the reproductive strategies of the species of fishes under investigation may be key to evaluating and understanding other possible roles of THs in the reproductive process.

ABBREVIATIONS CNS CRF E2 FSH FSH-R GnRH GPHa GSI GTH HPG HPT LH LH-R MAPK MT NCoR PTU rT3 RXR SMRT SS T T3 T4 TBG TH TR TRa1 TRE TRH TSH TSH-R TTR UV UVS

Central nervous system Corticotropin-releasing factor Estradiol Follicle-stimulating hormone Follicle-stimulating hormone receptor Gonadotropin-releasing hormone Glycoprotein hormone subunit a Gonadosomatic index Gonadotropin Hypothalamicepituitaryegonadal Hypothalamicepituitaryethyroid Luteinizing hormone Luteinizing hormone receptor Mitogen-activated protein kinase Methyltestosterone Nuclear receptor corepressor Propylthiouracil Reverse-triiodothyronine Retinoid X receptor Silencing mediator of retinoid and thyroid hormone receptor Somatostatin Testosterone Triiodothyronine Thyroxine Thyroxine-binding globulin Thyroid hormone Thyroid hormone receptor Thyroid hormone receptor isoform Thyroid response element Thyrotropin-releasing hormone Thyrotropin Thyrotropin receptor Transthyretin Ultraviolet Ultraviolet-sensitive

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Van der Geyten, S., Byamungu, N., Reyns, G. E., Kuhn, E. R., & Darras, V. M. (2005). Iodothyronine deiodinases and the control of plasma and tissue thyroid hormone levels in hyperthyroid tilapia (Oreochromis niloticus). J. Endocrinol., 184, 467e479. Van der Ven, L. T., Van den Brandhof, E. J., Vos, J. H., Power, D. M., & Wester, P. W. (2006). Effects of the antithyroid agent propylthiouracil in a partial life cycle assay with zebrafish. Environ. Sci. Technol., 40, 74e81. Van Overbeeke, & McBride, J. R. (1971). Hisotogical effects of 11-ketotestosterone, 17a-methyltestosterone, estradiol cypionate, and cortisol on the interrenal tissue, thyroid gland, and pituitary gland of gonadectomized sockeye salmon (Oncorhynchus nerka). J. Fish Res. Bd. Can., 28, 477e484. Vassart, G., & Dumont, J. E. (1992). The thyrotropin receptor and the regulation of thyrocyte function and growth. Endocr. Rev., 13, 596e611. Vassart, G., Pardo, L., & Costagliola, S. (2004). A molecular dissection of the glycoprotein hormone receptors. Trends Biochem. Sci., 29, 119e126. Veldhoen, K., Allison, W. T., Veldhoen, N., Anholt, B. R., Helbing, C. C., & Hawryshyn, C. W. (2006). Spatioetemporal characterization of retinal opsin gene expression during thyroid hormone-induced and natural development of rainbow trout. Vis. Neurosci., 23, 169e179. Vischer, H. F., & Bogerd, J. (2003). Cloning and functional characterization of a testicular TSH receptor cDNA from the African catfish (Clarias gariepinus). J. Mol. Endocrinol., 30, 227e238. Vtiger, R. T. (1987). Thyrotropin-releasing hormone and thyrotropin secretion. J. Lab. Clin. Med., 109, 327e335. Wabuke-Bunoti, M. A. N., & Firling, C. E. (1983). The prehatching development of the thyroid gland of the fathead minnow, Pimephales promelas (Rafinesque). Gen. Comp. Endocrinol., 49, 320e331. Wagner, M. S., Wajner, S. M., & Maia, A. L. (2009). Is there a role for thyroid hormone on spermatogenesis? Microsc. Res. Tech., 72, 796e808.


Wagner, M. S., Wajner, S. M., & Maia, A. L. (2008). The role of thyroid hormone in testicular development and function. J. Endocrinol., 199, 351e365. Wang, Y., Zhou, L., Yao, B., Li, C. J., & Gui, J. F. (2004). Differential expression of thyroid-stimulating hormone b subunit in gonads during sex reversal of orange-spotted and red-spotted groupers. Mol. Cell. Endocrinol., 220, 77e88. Weirich, R. T., Schwartz, H. L., & Oppenheimer, J. H. (1987). An analysis of the interrelationship of nuclear and plasma triiodothyronine in the sea lamprey, lake trout, and rat: evolutionary considerations. Endocrinology, 120, 664e677. Yamada, H., Hiriuchi, R., Gen, K., & Yamauchi, K. (1993). Involvement of four hormones in thyroxine deiodination in several tissues of immature yearling masu salmon, Oncorhynchus masou. Zool. Sci., 10, 587e596. Yamano, K., & Inui, Y. (1995). cDNA cloning of thyroid hormone receptor b for the Japanese flounder. Gen. Comp. Endocrinol., 99, 197e203. Yamano, K., Araki, K., Sekikawa, K., & Inui, Y. (1994). Cloning of thyroid hormone receptor genes expressed in metamorphosing flounder. Dev. Genet., 15, 378e382. Yamano, K., & Miwa, S. (1998). Differential gene expression of thyroid hormone receptor a and b in fish development. Gen. Comp. Endocrinol., 109, 75e85. Yen, P. M. (2001). Physiological and molecular basis of thyroid hormone action. Physiol. Rev., 81, 1097e1142. Yen, P. M., Ando, S., Feng, X., Liu, Y., Maruvada, P., & Xia, X. (2006). Thyroid hormone action at the cellular, genomic and target gene levels. Mol. Cell. Endocrinol., 246, 121e126. Yoshiura, Y., Sohn, Y. C., Munakata, A., Kobayashi, M., & Aida, K. (1999). Molecular cloning of the cDNA encoding the b subunit of thyrotropin and regulation of its gene expression by thyroid hormones in the goldfish, Carassius auratus. Fish Physiol. Biochem., 21, 201e210. Zoeller, R. T., Tan, S. W., & Tyl, R. W. (2007). General background on the hypothalamicepituitaryethyroid (HPT) axis. Crit. Rev. Toxicol., 37, 11e53.

Chapter 6

Stress and Reproduction Meghan L.M. Fuzzen, Nicholas J. Bernier and Glen Van Der Kraak University of Guelph, Guelph, ON, Canada

SUMMARY This chapter explores the interactions between the hypothalamice pituitaryeinterrenal (HPI) stress axis and the hypothalamice pituitaryegonadal (HPG) reproductive axis and their effects on reproductive processes in teleost fishes. We review the evidence that stress and activation of the HPI axis affect reproduction and do so through actions on the central nervous system, pituitary, and gonads, and production of hepatic vitellogenin (Vtg). Moreover, we describe how stress affects reproductive development at different life stages including embryonic and larval stages, at puberty, and as adults. Collectively, based on the studies conducted to date, it is not possible to provide a generalized model of how stress affects reproduction in teleosts. Rather, the influence of stress on reproduction depends on the type of stressor, its intensity and duration, the sex of the fish, its developmental stage, its nutritional status, and its reproductive strategy. Corticosteroids are primary mediators of the stress response, yet they exhibit both stimulatory and inhibitory influences on reproductive development, which adds to the complexity of defining the role of stress on reproduction in teleosts.

1. INTRODUCTION Stress is a common feature of life, and fishes, like all organisms, have evolved a suite of defense reactions to protect themselves against stimuli that pose a challenge to the maintenance of homeostatic equilibrium. A key component of the stress response in vertebrates is a reallocation of energy away from nonessential physiological functions, such as reproduction, and toward activities that contribute to the restoration of homeostasis. While this prioritizing of energy allocation is an integral component of allostasis, which represents part of the adaptive process for actively maintaining stability through change (McEwen & Wingfield, 2003), chronic inhibition of investment activities also can be maladaptive. The adverse consequences of chronic stress on reproduction have been well documented in several vertebrate groups including fishes (reviewed by Pankhurst & Van Der Kraak, 1997; Schreck, ContrerasSanchez, & Fitzpatrick, 2001; Milla, Wang, Madiki, Hormones and Reproduction of Vertebrates, Volume 1dFishes Copyright Ó 2011 Elsevier Inc. All rights reserved.

& Kestemont, 2009; Schreck, 2009; see also Volumes 2e5 in this series). In general, stressereproduction interactions are complex (Figure 6.1) as the various mediators of the stress response can impact on a broad range of reproductive functions and behaviors. This chapter focuses specifically on the interactions between the hypothalamicepituitaryeinterrenal (HPI) stress axis and the hypothalamicepituitaryegonadal (HPG) reproductive axis and the consequences of these interactions for reproductive processes. After a brief overview of the stress response and the regulation of reproductive functions in fishes, the chapter examines how stressors and the mediators of the stress response affect the key effectors of the HPG axis, including the hypothalamic gonadotropinreleasing hormones (GnRHs), the pituitary gonadotropins (GTHs), the gonadal sex steroids, and hepatic vitellogenin (Vtg). The chapter then reviews how stressors and stress hormones impact the development of the reproductive system and reproductive functions at the embryonic, larval, pubertal, and adult life stages. The effects of gender and reproduction on the activity of the HPI axis are discussed. Finally, we describe examples of fishes exhibiting an apparent resistance to stress during sexual maturation.

1.1. Effectors of the Stress Response The stress response in fishes is mediated by the HPI axis and the autonomic sympathetic-chromaffin cell axis (Wendelaar Bonga, 1997). In mammals, and presumably also in fishes, stress-sensitive brain circuits with both excitatory and inhibitory neurotransmitters coordinate the activation of both stress axes (Ulrich-Lai & Herman, 2009). At the hypothalamic level, among the multiple stimulatory and inhibitory hypophysiotropic factors that may be involved in regulating the HPI axis, corticotropin-releasing factor (CRF) from the nucleus preopticus (NPO) is considered the primary signal (Lederis, Fryer, Okawara, Schonrock, & Richter, 1994; Bernier, Flik, & Klaren, 2009). Depending on the reproductive status of a fish, urotensin I (UI), a CRFrelated peptide, and arginine vasotocin are additional 103

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FIGURE 6.1 Overview of the major neuroendocrine signals and interactions between the hypothalamicepituitarye interrenal (HPI) stress axis (in red) and the hypothalamicepituitaryegonadal (HPG) reproductive axis (in blue) in teleosts. Neurotransmitters regulate the activity of both axes. Whereas activation of the HPI axis by stressors results in the production of cortisol by the interrenals, the HPG axis stimulates the production of the sex steroids, estradiol (E2), testosterone (T), and 11ketotestosterone (11-KT). Estradiol also stimulates the production of vitellogenin by the liver. Solid black arrows indicate stimulation. Dashed black arrows indicate inhibition. Red arrows indicate potentials effects of cortisol and corticotropinreleasing factor (CRF) on the HPG axis. Blue arrows indicate potential effects of sex steroids on the HPI axis. 5-HT, serotonin; ACTH, corticotropin; Ct, corticotropes; DA, dopamine; FSH, follicle-stimulating hormone; GABA, g-aminobutyric acid; GnRH, gonadotropin-releasing hormone; Gt, gonadotropes; LH, luteinizing hormone; NA, noradrenaline. See color plate section.

hypophysiotropic factors that may play a significant role in the regulation of the HPI axis (Balment, Lu, Weybourne, & Warne, 2006; Westring et al., 2008). While CRF has multiple targets within the pituitary of teleosts, the hypophysiotropic role of CRF in the regulation of the hormone corticotropin (ACTH) secretion from the corticotropes is central to the workings of the HPI axis (Bernier et al., 2009). Circulating ACTH in turn is recognized as the principle regulator of corticosteroid synthesis in the interrenal cells of the head kidney and the stimulator of cortisol release during the acute phase of the stress response (Flik, Klaren, Van den Burg, Metz, & Huising, 2006; Aluru & Vijayan, 2008). Beyond its effects on reproduction, metabolism, growth, ionic balance, and the immune system, cortisol also can limit the magnitude and duration of the endocrine stress response in fishes via negative feedback effects on the gene expression of CRF in the NPO and the expression of the ACTH precursor molecule pro-opiomelanocortin (POMC) in the pituitary (Mommsen, Vijayan, & Moon, 1999; Norris & Hobbs, 2006; Bernier et al., 2009). In the sympathetic-chromaffin cell axis, the chromaffin cells of the head kidney release the catecholamines adrenaline and noradrenaline in response to activation by acetylcholine and non-neuronal pathways (Reid, Bernier, & Perry, 1998). While most stressors elicit the secretion of cortisol (Barton & Iwama, 1991; Barton, 2002; Norris & Hobbs, 2006), circulating catecholamine levels generally do not change in response to mild or moderate stress but

rise when fish experience acute life-threatening stressors (Perry & Bernier, 1999). Also, while noradrenaline of neuronal origin plays an important role in the neuroendocrine regulation of the HPG axis in fishes (Van Der Kraak, 2009), and sympathetic nerves are involved in the control of reproductive functions in mammals (Gerendai, Banczerowski, & Halasz, 2005), the role of catecholamines in the endocrine regulation of the HPG axis is poorly defined. Therefore, this review focuses on the interplay between the HPI and HPG axes, and the potential effects of humoral catecholamines on reproduction are not discussed.

1.2. Effectors of Reproductive Functions The HPG axis regulates reproductive functions in fishes (see Chapter 2, this volume) as in all vertebrates (see Volumes 2e5 in this series). At the brain and pituitary levels, multiple GnRH forms and GnRH receptors play central roles in coordinating reproductive endocrinology and behavior in teleosts (reviewed by Van Der Kraak, 2009). Gonadotropin-releasing hormone originating from the preoptic area (POA) induces GTH release from the pituitary, whereas the GnRH expressed outside the POA contributes to diverse neuromodulatory functions including the regulation of sexual behaviors (Soga, Ogawa, Millar, Sakuma, & Parhar, 2005; Kah et al., 2007). Gonadotropinreleasing hormone stimulates the secretion of the GTHsdfollicle-stimulating hormone (FSH) and luteinizing

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Stress and Reproduction

hormone (LH)dfrom the pituitary gonadotropes (Dickey & Swanson, 2000; Vacher, Mananos, Breton, Marmignon, & Saligaut, 2000; Ando & Urano, 2005; Aizen, Kasuto, Golan, Zakay, & Levavi-Sivan, 2007). Whereas GnRH is probably the most important stimulator of GTH release, the control of GTH secretion is multifactorial and involves several additional stimulatory and inhibitory peptides and neurotransmitters (Trudeau et al., 2000; Chang et al., 2009; Van Der Kraak, 2009). Multiple peptides and neurotransmitters also are involved in regulating the production of GnRH in the POA. One key regulatory factor of the HPG axis in teleosts is dopamine (DA) (Dufour et al., 2005). Dopamine potently inhibits basal and GnRH-stimulated LH secretion in most teleost species. There is also evidence that DA can inhibit both the release of pituitary FSH and POA GnRH. While FSH promotes gametogenesis in females by stimulating the production of 17b-estradiol (E2) and the incorporation of Vtg into developing oocytes, in males FSH stimulates Sertoli cell proliferation and testosterone (T) production, and maintains spermatogenesis. LH promotes final sexual maturation by stimulating gonadal steroidogenesis in both sexes, oocyte maturation and ovulation in females, and spermiation in males (Zmora, Kazeto, Kumar, Schulz, & Trant, 2007; Van Der Kraak, 2009). Beyond the role of GTHs, ovarian and testicular functions in teleosts are regulated by several other secondary hormones and growth factors (Van Der Kraak, Chang, & Janz, 1998; Van Der Kraak, 2009; see also Chapters 3 and 4, this volume). While these secondary endocrine and paracrine signals also may be affected by stressors, such interactions are beyond the scope of this review. In general, sex steroids participate in spermatogonial and oogonial proliferation and E2 also plays a key role in the synthesis of the egg yolk precursor Vtg in the liver. Finally, E2 and T exert positive and negative effects on the HPG axis. Whereas the negative feedback effects of sex steroids on GTH release are mediated by indirect effects on GnRH release via dopaminergic fibers, the positive feedback effects can be exerted either directly at the pituitary level or indirectly by effects on GnRH in the POA (Yaron et al., 2003; Levavi-Sivan, Biran, & Fireman, 2006; Van Der Kraak, 2009).

2. EFFECTS OF STRESS ON THE HYPOTHALAMICePITUITARYeGONADAL (HPG) AXIS The HPI and HPG axes can interact at multiple levels in teleosts (Figure 6.1). While components of the two axes are located in separate nuclei and organ structures, there is crosstalk between the two axes at all levels and in both directions. In this section we will discuss the effects of the HPI axis on the HPG axis; the reverse interaction will be discussed in Section 4.

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2.1. Effects of Stress on the Central Nervous System (CNS) There is limited information available with respect to the effects of stress or stress hormones on the hypothalamic components of the HPG axis in fishes. Adult male tilapia (Oreochromis niloticus) exposed to chronic social stress are reproductively inactive and have decreased whole brain mRNA levels of GnRH-I, the main hypophysiotropic form in most teleosts including tilapia, and GnRH-II, but not GnRH-III (Ogawa, Soga, Sakuma, & Parhar, 2003; see also Chapter 2, this volume). Similarly, juvenile carp (Cyprinus carpio) fed cortisol have lower whole brain levels of GnRH-III peptidedthe major hypophysiotropic form in carp, zebrafish (Danio rerio), rainbow trout (Oncorhynchus mykiss), and masu salmon (Oncorhynchus masou)dand an associated decrease in plasma LH content and delayed testicular development (Consten, Bogerd, Komen, Lambert, & Goos, 2001). It is likely that the changes in GnRH expression are a direct result of corticosteroid actions in the brain and hypothalamus. For example, in rainbow trout GnRH neurons in the caudal telencephalon/ anterior POA exhibit glucocorticoid receptor (GR) immunoreactivity (Teitsma et al., 1999). Further, glucocorticoid responsive elements (GREs) have been identified in the GnRH promoter of striped bass (Morone saxatilis) (Chow et al., 1998), tilapia (Farahmand, Rahman, Sohm, Hwang, & Maclean, 2003; Kitahashi, Sato, Sakuma, & Parhar, 2005), sea bream (Sparus sarba) (Hu et al., 2008), and zebrafish (Torgersen, Nourizadeh-Lillabadi, Husebye, & Alestrom, 2002). Overall, these results suggest that components of the stress axis may inhibit GnRH transcription and/or synthesis in fishes and more specifically that cortisol may directly regulate GnRH gene expression. In addition to glucocorticoids, various hypophysiotropic factors that regulate the HPI axis can affect the HPG axis and reproduction. For example, GnRH transcription in mammals is inhibited by CRF (Belsham & Lovejoy, 2005; Kinsey-Jones, Li, Bowe, Lightman, & O’Byrne, 2006; Keen-Rhinehart et al., 2009), arginine vasopressin (Tellam, Mohammad, & Lovejoy, 2000), UI, and sauvagine (Tellam et al., 1998). Although the specific mechanisms by which these neuropeptides affect GnRH are unknown, there is evidence suggesting that CRF may act directly on the transcription of GnRH through CRF receptors (CRF-Rs) or indirectly through a b-endorphin-mediated pathway (Tellam et al., 2000). Whether the CRF system or other hypophysiotropic regulators of the HPI axis affect the production of GnRH in fishes is an area for future research. Similarly, several neurotransmitters that contribute to the neurocircuitry of stress appear to be involved in the regulation of GnRH neurons and the control of reproduction (Dobson, Ghuman, Prabhakar, & Smith, 2003). For example, in the Atlantic croaker (Micropogonias

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undulates) a decrease in GnRH gene expression in the POA and an inhibition of reproduction have been observed following exposure to hypoxic conditions. This response was associated with decreased brain serotonin (5-HT) levels and the effects of hypoxic conditions on GnRH are reversible with the pharmacological restoration of 5-HT (Thomas, Rahman, Khan, & Kummer, 2007). Whether 5-HT also contributes to the regulation of the HPI axis in hypoxic Atlantic croaker is not known, but there is compelling evidence implicating the serotonergic system in the regulation of the stress response in fishes (S. Winberg, Y. Winberg, & Fernald, 1997; Ho¨glund, Balm, & Winberg, 2002). Although intricate relationships between the control of the HPI axis and GnRH neurons have been identified, it is still unclear how these interactions are mediated. Interestingly, there is now evidence that the kisspeptin (Kp) receptor (Kiss1r)-signaling system is an essential component in the regulation of GnRH transcription in mammals (Popa, Clifton, & Steiner, 2008; Roa, Aguilar, Dieguez, Pinilla, & Tena-Sempere, 2008) and fishes (Kitahashi, Ogawa, & Parhar, 2009). This system may play an important role in mediating the suppressive effects of stressors on reproduction. In rats, different stressors and intracerebroventricular injections of CRF can downregulate the gene expression of Kp and/or Kiss1r within both the POA and the arcuate nucleus of the brain (Kinsey-Jones et al., 2009). Although GnRH was not measured in this study, the suppression of Kiss1r signaling was associated with a reduction in LH release from the pituitary (Kinsey-Jones et al., 2009). The impact of stressors, CRF-related peptides, and cortisol on the Kp system in fishes is also an area for future research.

2.2. Effects of Stress at the Level of the Pituitary The impact of stress on LH secretion from the gonadotropes in fishes is equivocal. Chronic stress in rainbow trout (Bry & Zohar, 1980; Zohar, 1980) and acute stress in the white sucker (Catostomus commersoni) (Stacey, MacKenzie, Marchant, Kyle, & Peter, 1984) decrease plasma LH levels. Cortisol implants also reduce plasma LH levels in both male brown trout (Salmo trutta) and rainbow trout, and decrease pituitary LH content in male brown trout (Carragher, Sumpter, Pottinger, & Pickering, 1989). Pituitaries incubated in vitro from cortisol-fed juvenile carp release significantly less LH in response to GnRH stimulation than pituitaries from control fish (Consten, Lambert, & Goos, 2001). In contrast, plasma LH levels in cortisolfed carp are either similar to (Consten, Lambert, Komen, & Goos, 2002) or higher than (Consten et al., 2001c) the levels from control fish. Similarly, acute confinement of


male brown trout increase plasma LH levels (Pickering, Pottinger, Carragher, & Sumpter, 1987) and cortisol stimulates both LH production and LHb transcript levels in juvenile European eel pituitary cells (Anguilla anguilla) (Huang et al., 1999). These conflicting results are likely due to differences between species as well as differences in both the type and duration of the stressors. In general, the direct actions of cortisol on LH secretion from carp pituitary cells in vitro (Consten et al., 2001c), and the presence of GR immunoreactivity in the large majority of LH-like pituitary cells in rainbow trout (Teitsma et al., 1999), suggest a direct action of cortisol at the pituitary level in fishes. The FSH-like cells of the pituitary in rainbow trout also exhibit GR immunoreactivity (Teitsma et al., 1999) but, to our knowledge, only one study to date has measured the effect of cortisol on pituitary FSH gene expression in a fish and none have quantified the effects of stressors on plasma FSH levels. Immature common carp chronically fed cortisol-containing food pellets during the pubertal period have reduced pituitary FSHb mRNA levels (Consten et al., 2001a). In general, there appears to be a significant gap in our understanding of how stressors and different components of the HPI axis affect FSH transcription and translation.

2.3. Effects of Stress on Hepatic Vitellogenesis Since the initial reports showing that slow-release cortisolcontaining implants cause marked reductions in circulating Vtg levels in rainbow trout (Carragher et al., 1989), many studies using in-vivo and in-vitro approaches have demonstrated the inhibitory effects of corticosteroids on Vtg biosynthesis (e.g., Pellisero et al., 1993; Mori, Matsumoto, & Yokota, 1998; Teitsma et al., 1998; Lethimonier, Flouriot, Valotaire, Kah, & Ducouret, 2000; Berg, Modig, & Olsson, 2004; Berg, Westerlund, & Olsson, 2004). The actions of cortisol could be a result of direct effects in the liver and interference with the estrogendependent induction of Vtg production or through indirect mechanisms leading to a reduction in E2 levels. Studies with rainbow trout hepatocytes point to the former mechanism, as cortisol inhibits the expression of both the estrogen receptor (ER) and Vtg mRNA (Lethimonier et al., 2000; Leithmonier, Flouriot, Kah, & Ducouret, 2002). It seems that the ER may be the primary target as cortisol had a stronger inhibitory effect on ER mRNA expression compared to Vtg expression and did so with a higher sensitivity and a more rapid time course of action (Lethimonier et al., 2000). Other work showing that the promoter region of the ER contains GREs provides the functional basis for this effect (Teitsma et al., 1998). Further, activation of the GRE interferes with the actions of

Chapter | 6


a CCAAT/enhancer-binding protein b-like transcription factor that is involved in enhancing the transcription of the ER (Lethimonier et al., 2002). These studies are consistent with earlier work showing that cortisol implants reduce the number of hepatic ERs in rainbow trout (Pottinger & Pickering, 1990). It is becoming clear that there are species differences in the manner in which cortisol affects Vtg synthesis. For example, cortisol caused dose-dependent inhibition of estrogen-induced Vtg levels in the plasma of Arctic char (Salvelinus alpinus) but, unlike what was reported for the rainbow trout, cortisol had no effect on Vtg mRNA levels (Berg et al., 2004a). This suggests that cortisol may have post-transcriptional effects. In contrast to the inhibitory effects of cortisol on E2-induced Vtg induction, cortisol has been shown to potentiate the effects of E2 on the expression of the zona pellucida (eggshell) proteins in Arctic char (Berg et al., 2004b). These results indicate that Vtg and zona pellucida proteins in Arctic char are not regulated by the same mechanisms and this is reflected in a differential response to corticosteroids.

2.4. Effects of Stress on Gonadal Function The ovary and testis of teleosts synthesize corticosteroids including cortisol, 11-deoxycortisol, corticosterone, and 11-deoxycorticosterone (DOC) (Fostier, Jalabert, Billard, Breton, & Zohar, 1983; Kime, 1993; Milla et al., 2009). The ovary and testis also contain GRs and mineralocorticoid receptors (MRs) (Takeo, Hata, Segawa, Toyohara, & Yamashita, 1996; Sturm et al., 2005; Milla et al., 2008). Finally, there is evidence that the promoter region of steroidogenic enzymes such as aromatase contain GREs, providing a functional link for the actions of corticosteroids in regulating gonadal gene expression (Gardner, Anderson, Place, Dixon, & Elizur, 2005). Despite the numerous studies showing that the gonads produce and respond to corticosteroids, there is considerable uncertainty as to the physiological roles that corticosteroids play in gonadal tissues (Milla et al., 2009). Stress and/or cortisol reduce plasma sex steroid levels in a variety of teleost species (e.g., Pickering et al., 1987; Carragher et al., 1989; Jardine, Van Der Kraak, & Munkittrick, 1996; Clearwater & Pankhurst, 1997; Haddy & Pankhurst, 1999). Not surprisingly, many studies have evaluated the direct effects of corticosteroids on gonadal steroid biosynthesis. Although there was early work showing a direct suppressive effect of cortisol on E2 production by rainbow trout ovarian follicles (Carragher & Sumpter, 1990), subsequent studies found that cortisol was not directly associated with the inhibition of steroid production by goldfish (Carassius auratus), carp, New Zealand snapper (Pagrus auratus), or zebrafish

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ovarian follicles (Pankhurst, Van Der Kraak, & Peter, 1995; Alsop, Ings, & Vijayan, 2009). Given the large number of studies with ovarian tissues, it seems unlikely that cortisol has a direct inhibitory effect on ovarian steroidogenesis. There have been far fewer studies with the testis and these studies suggest that there are developmental differences in the responsiveness of the testis to corticosteroids. For example, Consten et al. (2001a) showed that the cortisol agonist dexamethasone blocked in-vitro production of 11-ketotestosterone (11-KT) by rainbow trout testicular tissue from pubertal animals (120 days post hatch (dph)) but had no effect in adolescent animals (greater than 165 dph). Recent studies point to another possible mechanism by which the stress axis may act directly at the level of the gonad to modulate steroid biosynthesis. This relates to work showing that the melanocortin 2 receptor (MC2R), which binds ACTH and is responsible for activating corticosteroid biosynthesis in the interrenal gland of fishes (Klovins et al., 2004; Aluru & Vijayan, 2008), is abundantly expressed in the ovary and testis of rainbow trout (Aluru & Vijayan, 2008). Corticotropin did not stimulate cortisol production in the zebrafish ovarian follicle (Alsop et al., 2009); however, ACTH did inhibit GTH-induced E2 production by zebrafish follicles incubated in vitro (Alsop et al., 2009). Corticosteroid levels in unstressed fishes are often highest around the time of spawning in females, which may reflect the energetic demands of this period. High levels of corticosteroids may also relate to the roles they play in oocyte maturation and ovulation. Corticosteroids promote meiotic maturation of full-grown ovarian follicles incubated in vitro (Figure 6.2a) (Goetz, 1983). It is unlikely that corticosteroids play a dominant role in this process in vivo, but they may promote the actions of progestins, which are the primary maturation-inducing steroids in fishes. Cortisol promotes hydration of the oocyte following oocyte maturation in rainbow trout (Figure 6.2b) (Milla, Jalabert, Rime, Prunet, & Bobe, 2006). Finally, a number of studies have shown that cortisol, 11-deoxycortisol, and DOC promote ovulation in vivo (Figure 6.2a) (Goetz & Theofan, 1979; Milla et al., 2009 for a review). There is evidence that DOC, which acts through the MR (Sturm et al., 2005), may play a fundamental role in reproduction in male teleosts. High levels of DOC have been detected around the time of spermiation in rainbow trout (Campbell, Fostier, Jalabert, & Truscott, 1980; Milla et al., 2008), and MRs are expressed in the testes and vasa deferentia in rainbow trout with levels of expression increasing during the initiation of spermiation (Milla et al., 2008). In-vitro incubations of rainbow trout testis pieces showed that DOC and cortisol cause a decrease in both basal and GTH-stimulated production of 17a,20b-P (Figure 6.2c) (Milla et al., 2008). This steroid is linked to various effects on spermiation in males including milt

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Collectively, these studies suggest that corticosteroids have effects on multiple processes associated with reproductive development and gamete maturation in fishes.

3. LIFE STAGE-SPECIFIC EFFECTS OF STRESS ON REPRODUCTION The effects of stress on reproduction often vary among species depending on the type and intensity of the stressor, and the timing of application. Although many studies have evaluated the effects of stress on gamete formation in adults, it is also important to consider the effects of stressful events at earlier life stages. The embryonic, larval, and juvenile stages are all critical time points in the life history and reproductive biology of fishes.

3.1. Impact of Stress During Embryonic and Larval Stages

FIGURE 6.2 The effects of corticosteroids on various aspects of gonadal function in fishes. (a) Effects of 11-deoxycortisol (11-DC) and 17a, 20bdihydroxyprogesterone (17a, 20b-P) (both at 31 ng/ml) on the proportion of yellow perch oocytes undergoing germinal vesicle breakdown (GVBD) and ovulation in-vitro. ), significantly different from the control. Redrawn from Goetz and Theofan (1979). (b) Effects of 17a, 20b-P (40 ng/ml), 11-deoxycorticosterone (DOC) (50 ng/ml), and cortisol (50 ng/ml) on the in-vitro hydration of rainbow trout ovarian follicles as assessed by the ratio of the initial wet mass of oocytes before (WMo) and after (WM) steroid treatment. IC, initial control; NC, negative control; ), significantly different from NC. Redrawn from Milla, Jalabert, Rime, Prunet, and Bobe (2006). (c) Effects of graded concentrations of DOC and cortisol (ng/ml) on basal and gonadotropin (GTH)-stimulated production of 17a, 20b-P by rainbow trout testis fragments. ), significantly different from the respective controls. Redrawn from Milla et al. (2008).

production, hydration, and the ionic composition of the seminal fluid, although there are species differences in the specific responses (Milla et al., 2008). DOC and 17a,20b-P together significantly reduce the spermatocrit, suggesting that they act together to increase the hydration of the milt. At this point the nature of the interactions between DOC and cortisol with 17a,20b-P are not well understood.

In teleosts, breeding females deposit cortisol into their eggs during gametogenesis. Utilization of cortisol begins shortly after fertilization, and cortisol is often completely depleted by the eyed stage of development (Auperin & Geslin, 2008). Stress during egg formation in females increases the quantity of maternal cortisol deposited into eggs (Stratholt, Donaldson, & Liley, 1997; McCormick, 1998). Although a few studies have found that an increase in maternal cortisol has no effect on embryonic growth or survival (Stratholt et al., 1997; Small, 2004), a larger number of studies show that elevated egg cortisol content has a negative impact on embryonic and larval development. For example, in tropical damselfish (Pomacentrus amboinensis), elevated maternal cortisol, either stress-induced or injected, is associated with a decrease in progeny size and increased variation in morphology (McCormick, 1998; 1999; McCormick & Nechaev, 2002; McCormick, 2006). High maternal cortisol levels are linked with low survivorship to the ‘eyed up’ stage in a study with masu salmon (Mingist, Kitani, Koide, & Ueda, 2007). Similar findings of decreased growth, increased mortality, and increased incidence of malformations are found in rainbow trout (Campbell, Pottinger, & Sumpter, 1992; 1994) and Atlantic salmon (Salmo salar) (Eriksen, Bakken, Espmark, Braastad, & Salte, 2006). Although the specific cellular mechanisms mediating the effects of elevated maternal cortisol on embryonic and larval development are not known, incubating eggs in cortisol induces similar effects (McCormick & Nechaev, 2002). There is evidence in mammals that exposure to elevated corticosteroids during development has adverse effects on gonadal development and reproductive behavior (Kaiser, Kruijver, Swaab, & Sachser, 2003; Ward et al., 2003; Meek, Schulz, & Keith, 2006; Piffer, Garcia, & Pereira, 2009).

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Exposure of developing embryos to various types of stressor also leads to effects similar to those seen with increasing maternal cortisol deposits in eggs, but it is unlikely that effects of early life stress are mediated by the same mechanism since the stress axis in most fishes is not functional until after hatching (Barry, Malison, Held, & Parrish, 1995; Jentoft, Held, Malison, & Barry, 2002; Pepels & Balm, 2004; Alsop & Vijayan, 2008). Embryos exposed to stressful conditions hatch earlier, are smaller in mass and length, and have an increased frequency of deformities (Rombough, 1988; Nguyen & Janssen, 2002; Ciuhandu, Stevens, & Wright, 2005; Eriksen et al., 2006; Hassell, Coutin, & Nugegoda, 2008). The negative impacts of maternal cortisol on hatchling size and weight can be augmented when combined with a stress applied during early development. For example, relative to Atlantic salmon embryos with elevated egg cortisol or to others exposed to hyperthermia, embryos with elevated egg cortisol that were also exposed to hyperthermia had a smaller fork length and body mass at hatch (Eriksen et al., 2006). The mechanisms mediating these effects are unknown and are an area for future research. Whether the size and weight reductions caused by early life exposure to stress have any impact on the reproductive capabilities of surviving progeny is still a matter of debate. There has been little research on effects of early life stress on subsequent sexual development and reproductive function of mature fishes. In zebrafish, exposure to hypoxic conditions (0.8 mg O2/liter) from five hours post fertilization to four months post fertilization resulted in a higher incidence of males (74%) relative to the normoxic controls (62%) (Shang, Yu, & Wu, 2006). The skewed sex ratio of zebrafish exposed to hypoxic conditions during development is also associated with decreased gene expression of several steroid biosynthetic enzymes and an increase in the testosterone/estrogen ratio (Shang et al., 2006). In the future, the use of rapidly developing fish models such as zebrafish and medaka (Oryzias latipes) should offer new opportunities to evaluate the effects of adverse rearing conditions on reproductive function and to study the mechanisms responsible for the possibly lifelong effects of maternal and early life stress.

3.2. Impacts of Stress on Puberty Prolonged cortisol treatment of common carp inhibits male pubertal development as measured by the first wave of spermatogenesis (Consten et al., 2001a; Consten, Keuning, Terlou, Lambert, & Goos, 2001b; Consten et al., 2001c; 2002a; 2002b). Juvenile carp around 60 dph were fed cortisol-treated food or received intraperitoneal implants of cortisol in cocoa butter. By approximately 90e100 dph, the cortisol-treated carp had a lower gonadosomatic index (GSI), impaired spermatogenesis, and lower plasma T and

11-KT levels as compared to controls. In-vitro studies confirmed that the latter effects were due to a decrease of the steroid production capacity of the testis (Consten et al., 2001b; 2002a; 2002b). Cortisol had modest effects on pituitary LH content and plasma LH levels, suggesting that the pituitary is not the primary site of the inhibitory effects of cortisol on testicular development (Consten et al., 2001a; 2001b). Rather, it seems likely that a decrease in androgen secretion may contribute to impaired maturation of gonadotropes. Interestingly, restoration of androgen levels in cortisol-treated carp did not result in testicular development similar to that of the control animals, suggesting that there also may be direct effects of cortisol on testicular development (Consten et al., 2002a). It is not known whether puberty in male fishes is uniquely sensitive to the effects of corticosteroids. However, at least for the carp, the effects of cortisol are most pronounced during pubertal development. Adolescent carp having completed the first wave of spermatogonial development and having been treated with cortisolcontaining food starting at 138 dph showed a diminished response to exogenous cortisol compared to pre-pubertal carp (Consten et al., 2002b). Cortisol administration promotes testicular development during the early stages of spermatogenesis, while inhibiting spermatogenesis during the mature phase in a freshwater fish, Notopterus notopterus (Shankar & Kulkarni, 2000). Studies of Japanese eel (Anguilla japonica) testis in an organ culture system show that cortisol induced DNA replication in spermatogonia and potentiated the actions of 11-KT on spermatogonial proliferation (Ozaki et al., 2006). Additionally, cortisol induces 11-KT production in eel testicular fragments (Ozaki et al., 2006). Collectively these studies suggest, perhaps unexpectedly, that in some species corticosteroids may play a positive role in spermatogonial proliferation. To date, there are no studies examining the effects of stress on the early stages of sexual development in female fish.

3.3. Impacts of Stress on Adults A number of studies have investigated the effects of stress and corticosteroids on reproductive development in sexually maturing fish. Most of these studies have shown that repeated stressful events delay ovulation. For example, redbelly tilapia (Tilapia zillii) fail to spawn in crowded holding tanks but the same fish will spawn soon after transfer to individual aquaria, coincident with increases in serum levels of E2 and T (Coward, Bormage, & Little, 1998). These studies showed that, as the duration of time individuals were held under crowded conditions increased, so did the incidence of ovarian atresia. This increased rate of atresia was associated with depressed steroid hormone levels (Coward et al., 1998). Increased atresia has been

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reported for vitellogenic follicles of the gurnard (Chelionichthys kumu) after confinement for up to 96 hours (Clearwater & Pankhurst, 1997). This observation contrasts with studies involving the striped trumpeter (Latris lineate), which showed that frequent handling of the broodstock resulted in a greater volume of eggs being produced by the handled fish (Morehead, Ritar, & Pankhurst, 2000). Similarly, adult female rainbow trout subjected to repeated acute handling stress during the later stages of reproductive development exhibited reduced egg size and a significant delay in ovulation, and males had lower sperm counts (Campbell et al., 1992). The same study also found significantly lower survival rates for progeny from stressed rainbow trout compared to progeny from unstressed control fish (Campbell et al., 1992). This is in sharp contrast to other studies showing that rainbow trout stressed during the period of final oocyte maturation and those that were stressed throughout vitellogenesis and oocyte maturation ovulated on average about two weeks earlier than controls (Contreras-Sańchez, Schreck, Fitzpatrick, & Pereira, 1998). This differs from the situation for rainbow trout stressed during the period of early vitellogenesis, which ovulated at the same time as controls. Absolute fecundity and fertilization were not significantly affected in any treatment group, but significant differences in relative fecundity were found. Stress applied early in vitellogenesis resulted in smaller eggs and swim-up fry (Contreras-Sańchez et al., 1998). Collectively these studies suggest that the response to stress varies markedly between species and depends on the time when the stress is applied and the nature of the stressor (Schreck et al., 2001). Corticosteroids also may play a role in regulating reproductive behavior in terms of mobilizing energy at times of mate selection, spawning, and nest defense. Whereas some studies have shown that corticosteroids are higher during these energetically expensive times, the relationship to reproduction is poorly understood (Knapp, 2003; Neff & Knapp, 2009). Several studies have investigated the effects of cortisol on reproductive function in adults. For example, sexually mature male brown trout treated with slow-release cortisol implants had significantly smaller testes, lower plasma levels of T, and reduced pituitary LH content compared to controls, although plasma 11-KT and LH were not different (Carragher et al., 1989). In contrast, testis size and plasma sex steroid levels in sexually maturing male rainbow trout were not affected by 36 days of cortisol treatment (Carragher et al., 1989). In other studies, sexually mature female brown trout implanted with cortisol for 18 days had smaller ovaries and reduced plasma levels of E2, T, and Vtg (Carragher et al., 1989). Similarly, sexually mature female Mozambique tilapia (Oreochromis mossambicus) implanted with cortisol-containing pellets for 18 days had


significantly depressed GSI, oocyte size, and serum T and E2 levels (Foo & Lam, 1993). Sex change is a common reproductive strategy in fishes (Devlin & Nagahama, 2002; see also Chapter 1, this volume) and corticosteroids have been hypothesized to play a role (Munday, Caley, & Jones, 1998; Perry & Grober, 2003; Frisch, Walker, McCormick, & Solomon-Lane, 2007). There are various ways in which this could operate but, for protogynous species (ones that first mature as a female and later change to a male), female fishes under suppressive male dominance and with high cortisol levels may be unable to change sex because the increased corticosteroids competitively inhibit the synthesis of 11-KT necessary for male development. Central to this issue are the overlapping actions of the enzymes 11b-hydroxylase and 11b-hydroxysteroid dehydrogenase, which are involved in the synthesis and deactivation of corticosteroids and also in the two-step synthesis of 11-KT from T (Kusakabe, Nakamura, & Young, 2003; Ozaki et al., 2006). It has been hypothesized that female fish (under suppressive male dominance) are unable to change sex because increased corticosteroid levels would inhibit 11-KT synthesis via substrate competition (Frisch et al., 2007). In an effort to test this hypothesis, female sandperch (Parapercis cylindrica) were treated with cortisol under conditions that were permissive to sex change. However, there was no effect of cortisol treatment on sex change or pattern of steroidogenesis, suggesting that increased corticosteroid has no effect on protogynous sex change in this species. (See Chapter 8, this volume for a discussion of hormonal regulation of sex change in fishes.)

4. EFFECTS OF SEX AND REPRODUCTION ON THE HYPOTHALAMICePITUITARYe INTERRENAL (HPI) AXIS There is considerable evidence that components of the HPG axis can affect the activity of the hypothalamicepituitarye adrenal (HPA) axis in mammals, the latter being the homolog of the HPI axis of fishes (Goel & Bale, 2009; Kudeilka, Hellhammer, & Wu¨st, 2009; Solomon & Herman, 2009). Although little research has been conducted on the effects of GnRHs or GTHs on the HPI axis, sex steroids do affect the latter. In general, the impact of sex steroids on the neuroendocrine stress response in fishes is of growing interest due to the large number of natural hormones and hormone mimics that are entering the aquatic environment (Arukwe, 2008; see also Chapter 13, this volume). To date, however, the results of studies on the impact of sex steroids on the stress response in fishes are equivocal and, although some studies suggest that E2 exacerbates and 11-KT attenuates cortisol secretion, others suggest the opposite.

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Noting larger increases in plasma ACTH and cortisol following a stressor in immature rainbow trout than in adult males, Pottinger, Balm, and Pickering (1995) first suggested that sex steroids may modulate the stress response in fishes and affect the activity of pituitary corticotropes. In a subsequent study, whereas sexually immature rainbow trout and immature brown trout of unknown sex given T and 11-KT implants had depressed stress-induced ACTH and cortisol levels, E2-implanted fish were characterized by an enhanced cortisol stress response (Pottinger, Carrick, Hughes, & Balm, 1996). Immature Atlantic salmon exposed to E2-containing water also had elevated plasma cortisol levels under both resting and stressed conditions (Lerner, Bjornsson, & McCormick, 2007). Interestingly, while the above studies suggest that E2 stimulates cortisol production in fishes, mature female rainbow trout with naturally high plasma E2 levels do not exhibit enhanced stress responsiveness (Pottinger et al., 1996). It has been suggested that high levels of T in both sexually maturing male and female fish may counteract the stimulatory effects of estrogens on the stress response (Pottinger & Carrick, 2000); however, this hypothesis has not been tested. In contrast, studies with juvenile gilthead seabream (Sparus aurata) (Teles, Pacheco, & Santos, 2005) and sea bass (Dicentrarchus labrax) (Teles, Pacheco, & Santos, 2006) have found that E2 injections or immersion depress plasma cortisol levels. In vitro, the rate of cortisol synthesis induced by pregnenolone and ACTH was higher in immature rainbow trout than in mature males, mature males given 11-KT implants, or immature females treated with 11-KT (Young, Thorarensen, & Davie, 1996). However, McQuillan, Lokman, and Young (2003) reported no effect of 11-KT on the rate of cortisol synthesis from the interrenals of juvenile or mature rainbow trout and Chinook salmon (Oncorhynchus tshawytscha), and an inhibitory effect of E2 on the ability of the head kidney to utilize pregnenolone for cortisol synthesis. McQuillan et al. (2003) also observed that immature and mature rainbow trout interrenals were insensitive to E2, and that mature female Chinook salmon and rainbow trout were more sensitive to ACTH stimulation than mature males. Similarly, Barry, Riebe, Parrish, and Malison (1997) observed no effect of either E2, T, or 11KT on basal cortisol secretion from incubated juvenile rainbow trout interrenals. Finally, Vijayan, Takemura, and Mommsen (2001) found no impact of E2 exposure on the cortisol levels of male Mozambique tilapia, and Carrera et al. (2007) saw no change in the cortisol levels of gilthead seabream injected with E2. While the reasons for these differing results are not known, differences in methodological approach (steroid dose, application, and duration) and species-specific differences, as observed between mammalian species (Young, Korszun, Figueiredo, Banks-Solomon, and Herman, 2008), are likely contributing factors. Despite the equivocal findings, and the

obvious need for a more reductionist approach in order to decipher the specific mechanisms involved, results from the above studies clearly implicate sex steroids as potential modulators of the HPI axis in teleosts.

5. REPRODUCTION AND RESISTANCE TO STRESS The chronic activation of the HPI axis that accompanies sexual maturation and spawning in Pacific salmon is paradoxical, as it appears to contradict the evidence presented in this chapter suggesting that the effectors of the HPI axis negatively impact reproductive functions in fishes. Characterized in numerous studies and species since the 1950s, the development of the gonads in Pacific salmon (genus Oncorhynchus) is accompanied by a sustained increase in plasma cortisol levels (for review see Dickhoff, 1989; Fagerlund, McBride, & Williams, 1995), hypertrophy and hyperplasia of pituitary corticotropes (Robertson & Wexler, 1962; Van Overbeeke & McBride, 1967), and elevated CRF and UI mRNA levels in the forebrain (Westring et al., 2008). Although resting plasma cortisol levels in immature Pacific salmon may not differ from those in other salmonids and typically lie below 10 ng ml 1, during sexual maturation and migration the levels can reach several hundred ng ml 1 and remain elevated for weeks to months (Robertson & Wexler, 1959; McBride, Fagerlund, Dye, & Bagshaw, 1986; Carruth et al., 2000; Westring et al., 2008). Further, although the demands of the spawning migration (e.g., changes in salinity, water temperature, water flow, migration distance) can influence the magnitude of the cortisol surge in maturing semelparous and anadromous Oncorhynchus species such as sockeye salmon (Oncorhynchus nerka) (Macdonald et al., 2000; Hinch, Cooke, Healey, & Farrell, 2006), HPI axis activation is also observed in maturing adults that are nonmigrating. For example, sexual maturation in captive sockeye salmon (Patterson, Macdonald, Hinch, Healey, & Farrell, 2004), landlocked kokanee salmon (O. nerka kennerlyi) (Carruth et al., 2000), and iteroparous migrating and nonmigrating rainbow trout (Robertson & Wexler 1959; Robertson et al., 1961) is characterized by a significant increase in HPI axis activity. These results suggest that the stimulation of the HPI axis in mature Pacific salmon is at least partly due to an endogenous programmed event and not solely the consequence of migration and spawning (Dickhoff, 1989; Carruth, Jones, & Norris, 2002). Interestingly, castration of sockeye and kokanee salmon reduces plasma cortisol levels, delays the activation of the HPI axis, and increases lifespan (Robertson, 1961; Robertson & Wexler, 1962; McBride & Van Overbeeke, 1969). Moreover, although the effects of sex

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steroids on the HPI axis are complex and equivocal (see Section 4), several studies show that estrogens and androgens can stimulate the HPI axis in gonadectomized Pacific salmon (Donaldson & Fagerlund, 1969; Fagerlund & Donaldson, 1969; Van Overbeeke & McBride, 1971). Therefore, while the physiological processes associated with senescence may contribute to the activation of the HPI axis in semelparous salmon, considerable evidence also suggests that sex steroids participate in this process. Although the specific physiological roles of the hyperactive HPI axis in sexually maturing Pacific salmon are unknown, the catabolic effects of cortisol presumably contribute to the mobilization of energy reserves, which fuels development of the gonads and the migration to the spawning grounds. Carruth et al. (2002) suggested that the increase in plasma cortisol associated with sexual maturation and migration may enhance the ability of Pacific salmon to recall the imprinted memory of the home-stream chemical composition. Although there is some evidence that the very high cortisol levels of salmon migrating through extreme conditions can be associated with a transient reduction in plasma sex steroid levels (Hinch et al., 2006), in general Pacific salmon successfully reproduce in the face of a prolonged surge in plasma cortisol. Thus, a marked and sustained increase in the HPI axis need not necessarily result in a suppression of the HPG axis and the energy-mobilizing properties of cortisol may take place without negatively affecting reproductive functions. Future studies on semelparous salmonids aimed at deciphering the cellular mechanisms by which the HPG axis overcomes the suppressive effects of the HPI axis during gonadal maturation could broaden our understanding of the interrelationships between the HPI and HPG axes in fishes.

6. CONCLUSIONS Stressors in fishes can impact all levels of the HPG axis and physiological processes that are essential for reproduction, from fertilization to spawning. While both central and peripheral effectors of the HPI axis likely play key roles in mediating the effects of stress on reproduction, to date the majority of studies have focused on the impact of cortisol and relatively little is known about the influence of the CRF system and ACTH on the regulation of reproductive functions. In general, although equivocal findings have been reported, cortisol mainly has a negative impact on early development, gonadal differentiation, puberty, gametogenesis, and sexual behavior. In contrast, cortisol may exert positive effects during oocyte maturation and ovulation, and in some species gonadal maturation proceeds during chronic HPI axis hyperactivity.


Despite a recognition that the regulation of GTH secretion and gonadal functions in teleosts are multifactorial processes, to date most of the studies investigating the impact of stress on reproduction have focused exclusively on the primary effectors of the HPG axis, i.e., GnRH, GTHs, and sex steroids. Future studies are therefore needed to take into consideration the potential impact of stressors on the various secondary endocrine and paracrine factors that are involved in regulating the growth, differentiation, and function of the reproductive system. Finally, with few exceptions, studies into the interactions between the HPI and HPG axes in fishes have been carried out on seasonal oviparous (egg-bearing) species. Fishes are characterized by an amazing diversity of additional reproductive tactics, including continuous oviparous batch spawning, various forms of viviparity (live-bearing), and hermaphroditism. Exploiting the diversity of life-history strategies and reproductive tactics among fish species may offer unique opportunities among vertebrates to decipher the intricate relationships between the stress and reproductive axes.

ABBREVIATIONS 11-KT 17a,20b-P 5-HT ACTH CRF CRF-R DA DOC dph E2 ER FSH GnRH GR GRE GSI GTH HPA HPG HPI Kiss1r Kp LH MC2R MR NPO POA POMC T UI Vtg

11-ketotestosterone 17a,20b-dihydroxyprogesterone Serotonin Corticotropin Corticotropin-releasing factor Corticotropin-releasing factor receptor Dopamine 11-deoxycorticosterone Days post hatch 17b-estradiol Estrogen receptor Follicle-stimulating hormone Gonadotropin-releasing hormone Glucocorticoid receptor Glucocorticoid response element Gonadosomatic index Gonadotropin Hypothalamicepituitaryeadrenal Hypothalamicepituitaryegonadal Hypothalamicepituitaryeinterrenal Kisspeptin receptor Kisspeptin Luteinizing hormone Melanocortin 2 receptor Mineralocorticoid receptor Nucleus preopticus Preoptic area Pro-opiomelanocortin Testosterone Urotensin I Vitellogenin

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Chapter | 6


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Piffer, R. C., Garcia, P. C., & Pereira, O. C. M. (2009). Adult partner preference and sexual behavior of male rats exposed prenatally to betamethasone. Physiol. Behav., 98, 163e167. Popa, S. M., Clifton, D. K., & Steiner, R. A. (2008). The role of kisspeptins and GPR54 in the neuroendocrine regulation of reproduction. Annu. Rev. Physiol., 70, 213e238. Pottinger, T. G., & Carrick, T. R. (2000). Contrasting seasonal modulation of the stress response in male and female rainbow trout. J. Fish Biol., 56, 667e675. Pottinger, T. G., & Pickering, A. D. (1990). The effect of cortisol administration on hepatic and plasma estradiol-binding capacity in immature female rainbow trout, Oncorhynchus mykiss. Gen. Comp. Endocrinol., 80, 264e273. Pottinger, T. G., Balm, P. H. M., & Pickering, A. D. (1995). Sexual maturity modifies the responsiveness of the pituitaryeinterrenal axis to stress in male rainbow trout. Gen. Comp. Endocrinol., 98, 311e320. Pottinger, T. G., Carrick, T. R., Hughes, S. E., & Balm, P. H. M. (1996). Testosterone, 11-ketotestosterone, and estradiol-17b modify baseline and stress-induced interrenal and corticotropic activity in trout. Gen. Comp. Endocrinol., 104, 284e295. Reid, S. G., Bernier, N. J., & Perry, S. F. (1998). The adrenergic stress response in fish: control of catecholamine storage and release. Comp. Biochem. Physiol., 120, 1e27. Roa, J., Aguilar, E., Dieguez, C., Pinilla, L., & Tena-Sempere, M. (2008). New frontiers in kisspeptin/GPR54 physiology as fundamental gatekeepers of reproductive function. Front. Neuroendocrinol, 29, 48e69. Robertson, O. H. (1961). Prolongation of the life span of Kokanee salmon (Oncorhynchus nerka kennerlyi) by castration before beginning of gonad development. Proc. Natl. Acad. Sci. USA, 47, 609e621. Robertson, O. H., & Wexler, B. C. (1959). Hyperplasia on the adrenal cortical tissue in Pacific salmon (genus Oncorhynchus) and rainbow trout (Salmo gairdnerii) accompanying sexual maturation and spawning. Endocrinology, 65, 225e238. Robertson, O. H., & Wexler, B. C. (1962). Histological changes in the organs and tissues of senile castrated kokanee salmon (Oncorhynchus nerka kennerlyi). Gen. Comp. Endocrinol., 2, 458e472. Robertson, O. H., Krupp, M. A., Thomas, S. F., Favour, C. B., Hane, S., & Wexler, B. C. (1961). Hyperadrenocorticism in spawning migratory and nonmigratory rainbow trout (Salmo gairdnerii); comparison with Pacific salmon (genus Oncorhynchus). Gen. Comp. Endocrinol., 1, 473e484. Rombough, P. J. (1988). Growth, aerobic metabolism, and dissolved oxygen requirements of embryos and alevins of steelhead, Salmo gairdneri. Can. J. Zool., 66, 651e660. Schreck, C. B. (2009). Stress and fish reproduction: the roles of allostasis and hormesis. Gen. Com. Endocrinol., 165, 549e556. Schreck, C. B., Contreras-Sanchez, W., & Fitzpatrick, M. S. (2001). Effects of stress on fish reproduction, gamete quality, and progeny. Aquaculture, 197, 3e24. Shang, E. H. H., Yu, R. M. K., & Wu, R. S. S. (2006). Hypoxia affects sex differentiation and development, leading to a male-dominated population in zebrafish (Danio rerio). Environ. Sci. Technol., 40, 3118e3122. Shankar, D. S., & Kulkarni, R. S. (2000). Effects of cortisol on testis of freshwater fish Notopterus notopterus (Pallas). Indian J. Exp. Biol., 38, 1227e1230. Small, B. C. (2004). Effect of dietary cortisol administration on growth and reproductive success of channel catfish. J. Fish Biol., 64, 589e596.

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Soga, T., Ogawa, S., Millar, R. P., Sakuma, Y., & Parhar, I. S. (2005). Localization of the three GnRH types and GnRH receptors in the brain of a cichlid fish: insights into their neuroendocrine and neuromodulator functions. J. Comp. Neurol., 487, 28e41. Solomon, M. B., & Herman, J. P. (2009). Sex differences in psychopathology: of gonads, adrenals and mental illness. Physiol. Behav., 97, 250e258. Stacey, N. E., MacKenzie, D. S., Marchant, T. A., Kyle, L., & Peter, R. E. (1984). Endocrine changes during natural spawning in the white sucker, Catostomus commersoni: I. Gonadotropin, growth hormone, and thyroid hormones. Gen. Comp. Endocrinol., 56, 333e348. Stratholt, M. L., Donaldson, E. M., & Liley, N. R. (1997). Stress induced elevation of plasma cortisol in adult female coho salmon (Oncorhynchus mykiss), is reflected in egg cortisol content, but does not appear to affect early development. Aquaculture, 158, 141e153. Sturm, A., Bury, N., Dengreville, L., Fagart, J., Flouriot, G., RafestinOblin, M. E., et al. (2005). 11-deoxycorticosterone is a potent agonist of the rainbow trout (Oncorhynchus mykiss) mineralocorticoid receptor. Endocrinology, 146, 47e55. Takeo, J., Hata, J. H., Segawa, C., Toyohara, H., & Yamashita, S. (1996). Fish glucocorticoid receptor with splicing variants in the DNA binding domain. FEBS Letters, 389, 244e248. Teitsma, C. A., Anglade, I., Lethimonier, C., Le Drean, G., Saligaut, D., Ducouret, B., et al. (1999). Glucocorticoid receptor immunoreactivity in neurons and pituitary cells implicated in reproductive functions in rainbow trout: a double immunohistochemical study. Biol. Reprod., 60, 642e650. Teitsma, C., Lethimonier, C., Tujague, M., Anglade, I., Saligaut, D., Bailhache, T., et al. (1998). Identification of potential sites of cortisol actions on the reproductive axis in rainbow trout. Comp. Biochem. Physiol. C: Pharmacol. Tox. Endocrinol., 119, 243e249. Teles, M., Pacheco, M., & Santos, M. A. (2005). Sparus aurata L. liver EROD and GST activities, plasma cortisol, lactate, glucose and erythrocytic nuclear anomalies following short-term exposure either to 17b-estradiol (E2) or E2 combined with 4-nonylphenol. Sci. Total Environ., 336, 57e69. Teles, M., Pacheco, M., & Santos, M. A. (2006). Biotransformation, stress and genotoxic effects of 17b-estradiol in juvenile sea bass (Dicentrarchus labrax L.). Environ. Int., 32, 470e477. Tellam, D. J., Mohammad, Y. N., & Lovejoy, D. A. (2000). Molecular integration of hypothalamoepituitaryeadrenal axis-related neurohormones on the GnRH neuron. Biochem. Cell Biol., 78, 205e216. Tellam, D. J., Perone, M. J., Dunn, I. C., Radovick, S., Brennand, J., Rivier, J. E., et al. (1998). Direct regulation of GnRH transcription by CRF-like peptides in an immortalized neuronal cell line. Neuroreport, 9, 3135e3140. Thomas, P., Rahman, M. S., Khan, I. A., & Kummer, J. A. (2007). Widespread endocrine disruption and reproductive impairment in an estuarine fish population exposed to seasonal hypoxia. Proc. R. Soc. B, 274, 2693e2702. Torgersen, J., Nourizadeh-Lillabadi, R., Husebye, H., & Alestrom, P. (2002). In silico and in situ characterization of the zebrafish (Danio rerio) gnrh3 (sGnRH) gene. BMC Genomics, 3, 25. Trudeau, V. L., Spanswick, D., Fraser, E. J., Lariviere, K., Crump, D., Chiu, S., et al. (2000). The role of amino acid neurotransmitters in the regulation of pituitary gonadotropin release in fish. Biochem. Cell Biol., 78, 241e259.

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Ulrich-Lai, Y. M., & Herman, J. P. (2009). Neural regulation of endocrine and autonomic stress responses. Nat. Rev. Neurosci., 10, 397e409. Vacher, C., Mananos, E. L., Breton, B., Marmignon, M. H., & Saligaut, C. (2000). Modulation of pituitary dopamine D1 or D2 receptors and secretion of follicle stimulating hormone and luteinizing hormone during the annual reproductive cycle of female rainbow trout. J. Neuroendocrinol., 12, 1219e1226. Van Der Kraak, G. (2009). The GnRH system and the neuroendocrine regulation of reproduction. In N. J. Bernier, G. Van Der Kraak, A. P. Farrell, & C. J. Brauner (Eds.), Fish Neuroendocrinology, Vol. 28 (pp. 115e149). Burlington, MA: Academic Press. Van Der Kraak, G., Chang, J. P., & Janz, D. M. (1998). Reproduction. In D. H. Evans (Ed.), The physiology of fishes (pp. 465e488). Boca Raton, FL: CRC Press. Van Overbeeke, A. P., & McBride, J. R. (1967). The pituitary gland of the sockeye (Oncorhynchus nerka) during sexual maturation and spawning. J. Fish Res. Board Can., 24, 1791e1810. Van Overbeeke, A. P., & McBride, J. R. (1971). Histological effects of 11ketotestosterone, 17a-methyltestosterone, estradiol, estradiol cypionate, and cortisol on the interrenal tissue, thyroid gland, and pituitary gland of gonadectomized sockeye salmon (Oncorhynchus nerka). J. Fish Res. Board Can., 28, 477e484. Vijayan, M. M., Takemura, A., & Mommsen, T. P. (2001). Estradiol impairs hyposmoregulatory capacity in the euryhaline tilapia. Oreochromis mossambicus. Am. J. Physiol. Regul. Integr. Comp. Physiol., 281, R1161e1168. Ward, I. L., Ward, O. B., Affuso, J. D., Long, W. D. I., French, J. A., & Hendricks, S. E. (2003). Fetal testosterone surge: specific modulations induced in male rats by maternal stress and/or alcohol consumption. Horm. Behav., 43, 531e539. Wendelaar Bonga, S. E. (1997). The stress response in fish. Physiol. Rev., 77, 591e625. Westring, C. G., Ando, H., Kitahashi, T., Bhandari, R. K., Ueda, H., Urano, A., et al. (2008). Seasonal changes in CRF-I and urotensin I transcript levels in masu salmon: correlation with cortisol secretion during spawning. Gen. Comp. Endocrinol., 155, 126e140. Winberg, S., Winberg, Y., & Fernald, R. D. (1997). Effect of social rank on brain monoaminergic activity in a cichlid fish. Brain Behav. Evol., 49, 230e236. Yaron, Z., Gur, G., Melamed, P., Rosenfeld, H., Elizur, A., & LevaviSivan, B. (2003). Regulation of fish gonadotropins. Int. Rev. Cytol., 225, 131e185. Young, E. A., Korszun, A., Figueiredo, H. F., Banks-Solomon, M., & Herman, J. P. (2008). Sex differences in HPA axis regulation. In J. B. Becker, K. J. Berkley, N. Geary, E. Hapson, J. P. Herman, & E. A. Young (Eds.), Sex Differences in the Brain: from Genes to Behavior (pp. 95e108). New York, NY: Oxford University Press. Young, G., Thorarensen, H., & Davie, P. S. (1996). 11-Ketotestosterone suppresses interrenal activity in rainbow trout (Oncorhynchus mykiss). Gen. Comp. Endocrinol., 103, 301e307. Zmora, N., Kazeto, Y., Kumar, R. S., Schulz, R. W., & Trant, J. M. (2007). Production of recombinant channel catfish (Ictalurus punctatus) FSH and LH in S2 Drosophila cell line and an indication of their different actions. J. Endocrinol., 194, 407e416. Zohar, Y. (1980). Dorsal aorta catheterization in rainbow trout (Salmo gairdneri). I. Its validity in the study of blood gonadotropin patterns. Reprod. Nutr. De´velop., 20, 1811e1823.


Chapter 7

Hormones and Sexual Behavior of Teleost Fishes David M. Gonçalves*, y and Rui F. Oliveira*, ** *

Instituto Superior de Psicologia Aplicada, Lisboa, Portugal, y Universidade do Algarve, Faro, Portugal, Instituto Gulbenkian de Cieˆncia, Oeiras, Portugal

**

SUMMARY Fishes are an excellent group for studying the mechanisms through which hormones modulate the expression of sexual behaviors in vertebrates. First, they have radiated virtually throughout all aquatic environments and this is reflected in an extraordinary diversity of mating systems and reproductive behaviors. Second, many species present a remarkable plasticity in their sexual displays, as exemplified by fishes that change sex or that adopt more than one reproductive tactic during their lifetime, and this plasticity seems to be mediated by hormones. Third, the fish neuroendocrine system is well conserved among vertebrates and the mechanisms of hormonal action in behavior are likely to share similarities with those of other vertebrates. We review the role of hormones and neuropeptides in the modulation of fish sexual displays. We also try to identify research areas in fish behavioral endocrinology that have the potential to be further developed (e.g., the role of hormones in the regulation of female sexual behavior) and some of the technological developments that promise to increase our knowledge in this field in the near future.

1. THEORETICAL CONSTRUCTS: APPETITIVE AND CONSUMMATORY PHASES In species with sexual reproduction, ‘sexual behavior’ may be generally defined as the set of behavioral acts directed towards the goal of producing offspring (see Munakata & Kobayashi, 2009 for an extended discussion). Classifying these behavioral acts into categories can be useful for descriptive purposes but in fishes the diversity of sexual displays challenges this classification. A widespread dichotomy applied in the context of sexual behavior is the appetitive/consummatory division and we begin by discussing its usefulness in fish behavioral endocrinology. The appetitive/consummatory division stemmed from the observations of early ethologists, who realized that, in a goal-directed behavioral sequence, behaviors are usually


more variable during the initial stages of the sequence and more stereotyped towards its end. To account for this variation, these researchers suggested that an initial appetitive phase, defined as the phase of searching towards the goal, can be distinguished from a final consummatory phase, defined as the stage when the goal is reached (Sherrington, 1906; Craig, 1917). Although this distinction is still widely applied in studies investigating the mechanisms of behavior, there is an ongoing debate on the usefulness of these terms. In a recent review, Sachs (2007) identified some problems in the current use of the appetitive/consummatory dichotomy. These include the difficulties in defining the boundary between the two phases and assigning a particular behavioral element to the appetitive or consummatory stage, and the historical link of these terms to early ethological models that have since been abandoned (e.g., the Lorenz hydraulic model (Lorenz, 1950)). In a reply, Ball and Balthazart (2008) argued in favor of the ongoing use of this terminology using two assumptions: (1) the appetitive/consummatory division is useful in mechanistic studies of animal behavior as it captures an important and real variation in behavioral displays during goal-seeking sequences. The authors agree that defining strict boundaries between the appetitive/ consummatory phases may sometimes be difficult and that some overlap should be taken into account when these terms are to be applied, but also suggest that this difficulty is common to other definitions in biology and not serious enough to overcome their usefulness. (2) The link between the appetitive/consummatory terminology and ancient ethological models is probably more historical than conceptual. The authors argue that theories underlying many scientific terms change over time without diminishing the present value of the terms. Likewise, the modern use of the appetitive/consummatory distinction is no longer conceptually linked to the early ethological models of motivation and reflects actual theories of behavior. 119

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When studying the role played by hormones in the regulation of fish sexual behavior, the use of the appetitive/ consummatory dichotomy raises two questions: (1) this distinction has been mainly applied to studies in birds and mammalsdcan fish sexual behavior also be partitioned into appetitive/consummatory phases? (2) If so, is this distinction useful when trying to understand the hormonal regulation of fish sexual behavior? The answer to the first question seems to be affirmative. Many fishes display a set of behaviors anticipatory to copulation/spawning that fits the definition of appetitive behaviors (i.e., behaviors displayed during the searchingeattracting phase for a sexual partner, typically more variable than consummatory behaviors). These include the establishment and defense of a breeding territory, the preparation of a spawning site (e.g., nest), and the expression of courtship displays. Consummatory (goal-reaching) behaviors include copulation in internal fertilizers or the spawning reflex in external fertilizers and are usually highly stereotyped. In some species the distinction and quantification of these behaviors is clear-cut, although the appetitive/consummatory division in the fish literature on sexual behavior is rarely employed. The answer to the second question, on whether this distinction is useful, is more debatable. The division of a continuous behavioral sequence directed towards the goal of mating into appetitive/consummatory phases is artificial and represents only one of many possibilities of partitioning sexual behavior sequences. One important point to consider in this regard is that the original division of sexual behavior into appetitive/consummatory phases made no mechanistic assumptions and merely tried to capture the natural variability of sexual behavior sequences. Some of the later criticism directed at the use of this division stems from the work of Frank Beach, who postulated the existence of two separate mechanisms for sexual processes: a sexual arousal (appetitive) mechanism and a copulatory and ejaculatory (consummatory) mechanism (Beach, 1942). However, the empirical evidence for these separate mechanisms is still not clear, and giving a mechanistic rather than simply descriptive value to the a-priori appetitive/consummatory distinction seems to be the origin of some of the later criticisms of the use of these terms (e.g., Sachs, 2007). As an example, an overview of the published literature on the neuroendocrine regulation of sexual behavior in rats, one of the most thoroughly investigated vertebrate models in this field, reveals some of these difficulties. In a review, Everitt (1990) suggests that the medial preoptic area (mPOA) of the hypothalamus is involved in the regulation of consummatory but not appetitive aspects of sexual behavior, whereas the ventral striatum regulates appetitive but not consummatory elements of sexual behavior. However, subsequent experiments have shown that the mPOA is also involved in appetitive aspects of


sexual behavior in this species (e.g., Paredes, Highland, & Karam, 1993) and the proposed difference in brain areas regulating appetitive and consummatory behaviors has been questioned (Hull et al., 1999; Paredes, 2003). Further, some studies have revealed that brain aromatization of testosterone (T) has activational effects in both appetitive and consummatory aspects of male rat sexual behavior (e.g., Roselli, Cross, Poonyagariyagorn, & Stadelman, 2003), while others have shown that aromatization of T is necessary to activate consummatory behaviors (copulation) but not appetitive behaviors (Vagell & McGinnis, 1997). Thus, the available data on the neuroendocrine regulation of sexual behavior in vertebrates is still scarce and partially contradictory, and only a very limited number of species have been investigated in detail. It is thus not possible to generalize on Frank Beach’s original proposal or to predict that the appetitive/consummatory division will be more useful than others when trying to understand general mechanisms of hormonal regulation of behavior. For now, it seems more reasonable to apply a-priori divisions of sexual behavioral sequences that have a descriptive rather than predictive value. In some cases, the appetitive/ consummatory division may be considered to be the most helpful for capturing the natural variation in behavioral sequences but other classifications will be more useful in other cases. This seems to be particularly true for fishes, which present wide variation in the patterns of sexual behavior.

2. PATTERNS OF SEXUAL BEHAVIOR Fishes have radiated virtually throughout all aquatic habitats and more than 31 500 species have so far been described (Froese & Pauly, 2009). This large number of species and the wide variation in the ecological conditions of their habitats have resulted in numerous morphological, physiological, and behavioral adaptations. This fact is reflected in their patterns of sexual behavior, which are extraordinarily diverse, ranging from species with nearly absent precopulation/prespawning behaviors to species with elaborate territorial, nest-building, and courtship displays. Although the patterns of sexual behavior in fishes have been reviewed elsewhere (e.g., Breder & Rosen, 1966; Thresher, 1984), a short overview of this diversity is presented here.

2.1. Defending and Preparing a Spawning Site The defense of reproductive resources seems to be the main cause of territoriality in fishes. In a comparative analysis using datasets for freshwater fishes of Canada (Scott & Crossman, 1979), marine fishes of eastern Canada (Scott & Scott, 1988),

Chapter | 7

Hormones and Sexual Behavior of Teleost Fishes

and tropical fishes (Thresher, 1984), Grant (1997) showed that territories for reproduction are more common than feeding territories. Further, in a comparative analysis between tropical and Atlantic temperate fish fauna, Almada, Henriques, and Gonçalves (1998) demonstrated that the occurrence of permanent territories decreases with latitude and this is likely to relate to the more seasonal reproductive patterns of temperate fish fauna. Territoriality associated with reproduction is uncommon in pelagic spawning, the main mode of reproduction in teleosts (Breder & Rosen, 1966; Balon, 1984; Thresher, 1984; Grant, 1997). However, in species with demersal eggs, reproduction often depends on the successful acquisition of a breeding territory. This is usually the first stage of the reproductive cycle but there are examples where territories are only established after mating and used not as spawning grounds but for the defense of eggs or juveniles. Breeding territoriality is associated with the defense of resources important for reproduction. These can be nesting structures or substrate, food, or shelters. Male territoriality is prevalent in fishes (e.g., Almada et al., 1998) but many examples of shared territoriality by a male and a female (e.g., Loiselle, 1977), cooperative territoriality (e.g., Taborsky, 1984), and even female territoriality (e.g., Tautz & Groot, 1975) have been described. Males can also defend females rather than territories and this has been documented both in pelagic and demersal spawners. For example, in the triggerfish Sufflamen chrysopterum, females compete over and defend territories (presumably for food or shelter) and males guard between one and three females. Males abandon the territories if females are experimentally removed, suggesting that males are guarding females rather than defending a territory (Seki et al., 2009). When territories are acquired prior to mating, two classes of reproductive behavior can be identified: agonistic displays used to acquire and defend reproductive territories and nest preparation/building behaviors. Within the same species, agonistic behaviors are displayed in many contexts and aggressive patterns exclusively associated with breeding territoriality seem to be rare or nonexistent. The enormous diversity of agonistic displays in fishes makes it difficult to search for ‘universal’ patterns. Examples of behavioral motor patterns included in the aggressive repertoire of teleosts include charging, butting, chasing, lateral and frontal displays, opercular flare, fin spreading, circling, pendeling (back and forth swimming between two males in frontal position), and mouth-to-mouth fighting (Figure 7.1(a)). Submissive behaviors include fleeing, head-up positions, and body tilting (Figure 7.1(b)). Signals in other sensorial modalities (electrical, acoustic, chemical) may also be used during aggressive interactions. As an example, in the Mozambique tilapia (Oreochromis mossambicus) territorial males store urine and release it in

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FIGURE 7.1 Examples of agonistic displays in fish. (a) Mouth-to-mouth fighting in the Mozambique tilapia (Oreochromis mossambicus), a high-intensity aggressive display. (b) Dorsal presentation in the blenny Parablennius sanguinolentus parvicornis, a submissive posture. Reproduced from Santos and Barreiros (1993).

pulses during agonistic interactions with other males. As the olfactory potency of the urine correlates with the male’s social status, males may be signaling their status to other males using chemical cues in the urine (Barata, Hubbard, Almeida, Miranda, & Cana´rio, 2007; see also Chapter 9, this volume). Fishes that secure breeding territories also may perform behaviors associated with the preparation of spawning or nesting structures. This is more frequent in species with parental care, and the complexity of the nesting structures seems to correlate with the extent of parental care (Potts, 1984). Again, these behaviors are highly diverse among species. In its simplest form, a depression in the substrate where eggs are deposited is dug usually by the male using vigorous tail undulations (lateral or vertical) and/or pushing and mouth digging (Figure 7.2(a)). In some species males additionally build sandcastles or sand-scrape structures as display sites to attract females, and nest features can be assessed by females to select a mating partner (e.g., Fryer & Iles, 1972; Tweddle et al., 1998; Barber, Nairn, & Huntingford, 2001). Parental care of the substrate-deposited eggs may be performed by the male (e.g., Spondyliosoma cantharus (Wilson, 1958)), the female (e.g., Radulinopsis taranetzi (Abe & Munehara, 2005)), or both (e.g., Tilapia zilli (Loiselle, 1977)). In mouth-brooders one or both parents take the eggs (or fry in the case of late mouth-brooders) into the mouth and incubate them for a variable period of time (e.g., Balshine-Earn & Earn, 1998).

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FIGURE 7.2 Examples of nest-building in fishes. (a) Mouth-digging in Oreochromis mossambicus. (b) A female three-spined stickleback (Gasterosteus aculeatus) inside an algal nest. Reproduced from Pelkwijk and Tinbergen (1937). (c) A male fighting fish (Betta splendens) close to its bubble nest.

Natural crevices or holes can also be used as nesting sites, as e.g. in some gobies and blennies (Breder & Rosen, 1966). These natural cavities may receive no or minimal modifications, which include digging underneath a stone or a shell to increase the available space, cleaning the area in front of the nest entrance, and building structures in front or on top of the nest. Typically, the eggs are deposited inside the nest and the male provides parental care of the eggs or juveniles (e.g., Breder & Rosen, 1966; Thresher, 1984). More complex nesting behaviors also have been described in fishes. The three-spined stickleback (Gasterosteus aculeatus) digs a pit in the substrate and then collects with the mouth algal pieces that are bound in a tunnel shape with spiggin, a glycoprotein produced by the kidney (Wootton, 1976) (Figure 7.2(b)). The male leads the female into this tunnel and eggs are laid and fertilized inside the algal nest. After receiving eggs from several females, the male switches into the parental phase and protects the eggs until hatching (Kraak, Bakker, & Mundwiler, 1999). Another species that builds algal nests is the corkwing wrasse (Symphodus melops). Males gather algal pieces of several species and deposit them along a crevice. Softer algae are deposited in the deepest part of the nest, where eggs will be laid, presumably because they offer more suitable conditions for egg development. Towards the outer face of the nest harder algae are deposited, forming a network that holds the nest together. After each egg laying, the male expels the female from the nest and covers the egg mass with soft algae, resuming courtship and receiving more eggs afterwards (for details see Potts, 1974; 1984). In a final example, freshwater species such as the fighting fish Betta splendens build a bubble nest with a mixture of mucus and water and defend a territory around it (Figure 7.2(c)). Males attract females into the nest and after egg laying the male assumes parental care of the eggs and juveniles (Forselius, 1957). The examples described here illustrate the diversity of territorial and nest-building behaviors in fishes. After establishing a territory and preparing the breeding ground, fishes have to attract and select a mating partner.

2.2. Finding and Choosing a Mate: Species Identification, Sexual Discrimination, and Mate Choice Reproduction depends on the correct identification of the species and gender of potential partners. As we shall see, this is not always an easy task. Let us illustrate the mechanisms of species and gender recognition in fishes with two examples: electric fish and cichlids. The Gymnotiformes from South America and the Mormyriformes from Africa produce weak electric signals that are used for electrolocation and in intraspecific communication (Zakon & Smith, 2002). These signals are produced in electric organs located in the tails and received by specialized electroreceptors mainly located in the midline of the fishes (Zakon & Smith, 2002). More than 20 species producing similar signals may occur in sympatry in turbid rivers (Hopkins, 1980) and this presents a challenge for species and gender recognition. The first question is whether the electric organ discharge (EOD) signal has the potential of being used for these purposes. The EOD signal is highly stereotyped and species-specific, and its properties depend on the anatomy and physiology of the electric organ (e.g., Kramer, Kirschbaum, & Markl, 1981; Hopkins, 1988; 1999). In most species there are also sex differences in the EOD signal properties and even individual ‘EOD signatures’ (Hopkins, 1981; 1988; Crawford, 1992; McGregor & Westby, 1992; Friedman & Hopkins, 1996). Thus, the intrinsic differences in the EOD signal properties between and within species have the potential to be used in species, gender, and even individual recognition. This has been confirmed to some extent by a number of studies. For example, females of the mormyrid Campylomormyrus compressirostris have been found to prefer the EOD signal of a conspecific male over an EOD signal produced by males of the closely related sympatric Campylomormyrus rhynchophorus. However, females failed to discriminate the EOD signal of conspecific males from that of another sympatric species, Campylomormyrus tamandua, the EOD signal of which is more similar to

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conspecific males (Feulner, Plath, Engelmann, Kirschbaum, & Tiedemann, 2008). Using synthetic playbacks, Hopkins and Bass (1981) showed in a mormyrid species that males can recognize and respond to the signal from conspecific females based only on the temporal properties of the waveform. Further, males of another mormyrid (Polimyrus isidori) have been found to discriminate between the electrical signals of conspecific males and females, but in this species the sequence of pulse intervals and not the EOD waveform was used for discrimination (Crawford, 1991). Taken together, the results from these and many other studies (see Carlson, 2002 for a review) show that the EOD signal is used by electric fish to recognize the species and the gender of the sender, a fundamental first step in mating. Cichlids have rapidly radiated into a large number of diverse species (e.g., Johnson et al., 1996). The coexistence of a large number of species in the same area, often very similarde.g., more than 500 species of cichlid have been described for Lake Malawi (Martens, 1997)dsuggests fine mechanisms must be present for species and gender recognition. Males are usually more colorful than females and color differences are more marked between sexually active males of different species than between females or nonreproductive males (Fryer & Iles, 1972). In a study on the Pseudotropheus zebra species complex, females preferred conspecific males over heterospecific males from three closely related species, and, when presented only with the three heterospecific males, females preferred to associate with the male with the most similar color pattern to the conspecific (Couldridge & Alexander, 2002). In another study, differences in color patterns between males of two closely related haplochromine cichlids were manipulated using light conditions, and females were shown to prefer conspecific males when color differences were visible. However, when color differences were masked by light conditions, females of both species did not discriminate between conspecific or heterospecific males (Seehausen & Alphen, 1998). This suggests that color patterns are used by female cichlids for recognizing conspecific mating partners (see also Kidd, Danley, & Kocher, 2006; Stelkens et al., 2008). The males’ ability to correctly identify conspecific females based on visual cues alone seems to be poorer than the females’ ability to visually discriminate conspecific males (Knight & Turner, 1999), corroborating the predicted stronger selective pressure for correct species and gender identification in females. It should be noted that other studies suggest that both chemical (Plenderleith, Van Oosterhout, Robinson, & Turner, 2005) and acoustic (Amorim, Knight, Stratoudakis, & Turner, 2004; Amorim, Simo˜es, Fonseca, & Turner, 2008) cues also may be important in cichlids for species and gender discrimination, and most likely interspecific mate recognition is mediated by multimodal signals. The above examples show how

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long-range cues or signals detected in several sensorial modalities are used by animals to find and approach appropriate mates. Additionally, animals usually display some form of mate choice and the most extravagant morphologic structures and elaborate behaviors in nature seem to be the consequence of sexual selection (Darwin, 1871). Sexual selection is predicted to be stronger in males and mating partner selectivity is expected to be stronger in females (Trivers, 1972; Andersson, 1984). Fishes follow this pattern and males generally exhibit the most elaborate courtship signals and are more ornamented than females. However, many exceptions exist. For example, sexual dimorphism and dichromatism is absent in many species, notoriously in most pelagic spawners (Breder & Rosen, 1966; Thresher, 1984), and even when it occurs both females and males may apparently accept any mate, as long as it is recognized as a conspecific of the correct sex (e.g., Goulet & Green, 1988). Other exceptions discussed below are sex-rolereversed species in which females compete for mates more intensely than males do for females and may be the more ornamented sex (e.g., Berglund, Widemo, & Rosenqvist, 2005). It should be noted that, in species with traditional sex-roles, female traits are also under sexual selection by male choice, although the intensity of sexual selection is expected to be stronger on male traits as male potential reproductive rates are higher than those of females (Amundsen & Forsgren, 2001). The rule in species with sexual dimorphism is nevertheless for males to be more ornamented than females and to take the initiative in courtship (for a review on the function of male ornaments in animals see Berglund, Bisazza, & Pilastro, 1996). In many fishes, males in reproductive condition react to the presence of females with conspicuous courtship displays that may involve stereotyped motor patterns, color changes, pheromone release, and vocalizations or the production of electrical signals. The properties of these signals may correlate with male traits, and females can use this information to select mates and gain direct or indirect benefits. For instance, in the bicolor damselfish (Stegastes partitus), females select males that court at higher rates. This trait, courtship rate, correlates with egg survival, probably because males that court more are in better condition and provide more suitable parental care to the eggs (Knapp & Kovach, 1991). Thus, courtship rate can be considered an honest indicator of male condition and females gain direct (egg survival) or indirect (if variation in male condition has a genetic influence) benefits by selecting high-frequency-courting males. The same traits used for species/gender discrimination also may be under sexual selection by female choice. In a mormyrid, the sexually dimorphic EOD signal is longer in high-status males than in low-status males (Carlson, Hopkins, & Thomas, 2000). In the pintail knifefish Brachyhypopomus

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pinnicaudatus, females prefer larger males over smaller males, and the EOD amplitude and duration are correlated with male body size (Curtis & Stoddard, 2003). Thus, there is the potential for females to use the properties of the EOD signals for both species and gender identification as well as for mate choice. Feulner, Kirschbaum, Mamonekene, Ketmaier, and Tiedemann (2007) suggest for mormyrids of the genus Campylomormyrus inhabiting the Congo River that divergent selection acting on the feeding apparatus leads to changes in the EOD signal and to speciation by sexual selection and assortative mating based on the characteristics of this signal. In cichlids, variation in female preference for male colors seems to drive male color polymorphism and sympatric speciation by divergent sensory drive (Dominey, 1984; Seehausen et al., 2008; Seehausen, Van Alphen, & Witte, 1997). The species-specific traits used by females to identify conspecific males have therefore evolved by disruptive sexual selection driven by female mate choice. Similarly, individual variation in male preference for female color morphs has been also shown in the cichlid Neochromis omnicaeruleus, suggesting a potential for speciation driven by male mating preferences on female coloration (Pierotti, Martin-Fernandez, & Seehausen, 2009). Again, searching for general patterns in courtship displays in fishes is difficult due to their enormous interspecific variability. Some examples of male motor patterns displayed during courtship and that illustrate this variability are the sigmoid posture of poeciliids, the zigzag dance of sticklebacks, the lateral body quivering of zebrafish, and the circling and figure-eight swimming of blennioids (Figure 7.3).

2.3. Internal and External Fertilization In teleost fishes, external fertilization is predominant over internal fertilization and is considered to be the ancestral


condition. In species with external fertilization, sperm and egg release may be synchronous or asynchronous. Synchronous spawning seems to be predominant in pelagic spawners. When external fertilization takes place in the water-column between paired fish, the male and the female may assume positions that bring their genital pores close and synchronize the release of eggs and sperm (Breder & Rosen, 1966). A common strategy in group spawning species is for several males to follow a gravid female and compete for a privileged position close to her. When the female assumes a spawning posture, males closer to the female try to release sperm synchronously with egg release (Domeier & Colin, 1997; Kiflawi, Mazeroll, & Goulet, 1998). In asynchronous spawning species, males may release sperm prior to or after egg laying. For instance, males of some gobies perform upside-down movements inside the nest, during which they attach sperm trails to the nest walls prior to oviposition by the female (e.g., Marconato, Rasotto, & Mazzoldi, 1996; Ota, Marchesan, & Ferrero, 1996). Sperm release after oviposition occurs, for instance, in haplochromine cichlids. Females deposit the eggs in the male’s nesting pit and immediately afterwards collect them with the mouth. Attracted by the anal fin egg-spots characteristic of haplochromine male cichlids, females approach the male’s genital region and try to pick up these egg mimics. Males then release sperm and fertilization takes place inside the female’s mouth (Fryer & Iles, 1972; see also Salzburger, Braasch, & Meyer, 2007). In species with internal fertilization, females may cooperate with males or not. In the first case, females expose the genital pore to males and facilitate intromission of the male’s copulatory organ. In the second case, a male chases a female while trying to insert the copulatory organ into the female’s genital pore. Detailed descriptions of these behaviors for the guppy Poecilia reticulata can be found in Liley (1966) and Houde (1997).

FIGURE 7.3 Examples of sexual displays in fishes. (a) The zig-zag dance of Gasterosteus aculeatus. Redrawn from Tinbergen (1951). (b) A male Parablennius sanguinolentus parvicornis ‘circling’ a female from his nest. Reproduced from Santos and Barreiros (1993).

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Extraordinary exceptions to the general pattern of egg fertilization in animals are seahorses and pipefishes (Syngnathidae). In some genera, males have brood pouches and females transfer the eggs by inserting their ovipositor into the male’s brood pouch. Egg fertilization takes place inside the male’s pouch, and the male incubates the eggs until hatching (Fiedler, 1954). Motor patterns associated with gamete release are less variable than behaviors preceding this step. Sperm ejaculation is usually accompanied by stereotyped motor patterns consisting of high-frequency body quivering and are often similar between species with internal and external fertilization (Breder & Rosen, 1966). Egg laying may also be accompanied by quivering, whereas in live bearers parturition consists of lower-frequency contractions of the abdominal cavity, or of the pouch walls in the case of male syngnathids (Breder & Rosen, 1966).

2.4. Sex-role Reversal It is clear from the above examples that in most species females assume a passive role during the prespawning/ precopulation phase and have poorly elaborated courtship displays. For this reason, studies on female reproductive behavior in fishes have been mainly focused on consummatory aspects, although a few species, notoriously the goldfish, have been investigated in more detail. This lack of knowledge is somewhat surprising as the wide variation in fish behavior offers an opportunity to overcome this gap. Species with sex-role reversal, where females take the initiative in courtship and display courtship behaviors more frequently than males, are particularly well-suited to studying the proximate mechanisms regulating female reproductive displays. Sex-role reversal occurs when the males’ potential reproductive rates are lower than those of females. In these cases females will compete more intensely over mates, and males are expected to be more selective than females (Clutton-Brock & Vincent, 1991). In teleost fishes, the paradigmatic example of sex-role reversal are syngnathids (for a review see Berglund & Rosenqvist, 2003). Male seahorses and pipefishes carry the eggs until hatching and thus invest more heavily than females in parental care. However, a strong paternal investment seems to be a necessary but not sufficient prerequisite for sex-role reversal, and in syngnathids sex-roles depend on the mating system. In monogamous species, such as seahorses, the female opportunities for additional mating are reduced and the sex-roles are conventional, while in polygamous species, such as pipefish, the sex-roles are reversed (Vincent, 1992; 1994). Two well-studied syngnathids with sexrole reversal are Nerophis ophidion and Syngnathus typhle. The adult female-to-male ratio for these species in nature is approximately 1 : 1, and females produce more eggs than

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one male brood pouch can accommodate (Berglund, Rosenqvist, & Svensson, 1989; Berglund & Rosenqvist, 1990). This results in a female-biased operational sex-ratio with more females in reproductive condition than males, as suggested by the observation that most males have a full brood pouch at the peak of the breeding season (Berglund & Rosenqvist, 1993). As a consequence, females compete for mates more strongly than males, and males are more selective than females, preferring to mate with larger females (Berglund, Rosenqvist, & Svensson, 1986; Rosenqvist, 1990). Females are larger and, particularly in N. ophidion, more ornamented than males. Female ornamentation consists of a permanent blue coloring along the flanks and the development of a ventral skin fold during the breeding season. Females display towards males, often in lek-like aggregations, by swimming up and down above the eelgrass. Females compete amongst themselves for access to males and may force other females off a male. Female S. typhle use these ornaments both for male attraction and to intimidate other females (Berglund & Rosenqvist, 2009). After selecting a female, a male will dance with her above the eelgrass with wriggling and shaking movements, and this dance may precede copulation (Fiedler, 1954; Vincent, Berglund, & Ahnesjo¨, 1995; Berglund & Rosenqvist, 2003). Within the same species, the potential reproductive rates of males and females may also vary between populations and throughout the breeding season in the same population. In the two-spotted goby (Gobiusculus flavescens), at the beginning of the breeding season the adult female to male ratio is approximately 1 : 1 and sex-roles are conventional. However, as the season progresses the proportion of males sharply decreases, probably due to a higher male than female mortality rate. This is accompanied by a shift in sex-roles with females competing aggressively for the few available males and taking the initiative in courtship (Forsgren, Amundsen, Borg, & Bjelvenmark, 2004). In the peacock blenny Salaria pavo, two populations occurring in different ecological scenarios also differ in sex-roles. A population at the Adriatic Sea occurs in a rocky shore area where nest availability is high. Males reproduce in rock crevices and actively defend an area around the nest. Sex-roles are conventional, with males taking the initiative in courtship and performing more courtship displays than females (Saraiva, 2009). In contrast, in southern Portugal, the species inhabits a coastal lagoon where appropriate nesting substrate is scarce. Most males use artificial substrates such as brick holes for nests and do not defend any area around the nest. The scarcity of nests biases the operational sex ratio towards females and in this population females take the initiative and display more courtship behavior than males (Almada, Gonçalves, Oliveira, & Santos, 1995; Saraiva, 2009).

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2.5. Alternative Reproductive Tactics In some species, individuals of the same sex, usually males, may adopt alternative tactics to reproduce. Large ‘bourgeois’ males generally defend reproductive resources such as females or nests whereas smaller males parasitize the investment of bourgeois males (Taborsky, 1997). Alternative reproductive tactics (ARTs) can be categorized as fixed, when individuals adopt one of the tactics for their entire lifetime, or plastic, when individuals change tactics during their lifetime (Moore, 1991; Brockmann, 2001). Plastic tactics can further be classified as irreversible, when individuals switch from one tactic to another at a particular moment in their lifetime, and reversible, when individuals change back and forth between tactics (Moore, 1991; Moore, Hews, & Knapp, 1998; Brockmann, 2001). Teleost fishes are the vertebrate group with the highest incidence of male alternative reproductive tactics (Taborsky, 1994; 2008) and the most common pattern is for males to start reproducing parasitically and then irreversibly switch into the bourgeois tactic (reviewed in Oliveira, 2006). Traits related to female attraction and monopolization of reproductive resources are selected in bourgeois males, whereas traits prevail in parasitic males that increase the probability of stealing fertilizations. This may result in polyphenisms within the same sex and often parasitic and bourgeois males differ markedly in behavior, morphology, and physiology. The reproductive behavior of bourgeois males follows the general pattern above described for male teleosts, with the access to reproductive resources depending on territorial, nesting, and/or courtship displays. Parasitic males access reproductive resources and fertilize eggs in different ways. These include trying to approach nests or females inconspicuously, darting by fast swims into nests, forcing copulations with females in species with internal fertilization, or relying on a female-like appearance and behavior to approach nesting males (Taborsky, 1999; 2008). From a behavioral neuroendocrine perspective, species in which parasitic males reproduce by female-mimicry are particularly interesting, as male- and female-like traits coexist in the same individual. These fishes have a femalelike appearance and may display female-like behaviors, and simultaneously need to develop functional testes and other male traits. Thus, the neuroendocrine characterization of these species may provide insights into the role played by the endocrine system in the regulation of sexually dimorphic morphologic and behavioral traits. A species for which female-mimicry has been studied in some detail is the peacock blenny. In this species sexual dimorphism is intense, with bourgeois males being larger than females and having a set of well-developed secondary sexual characteristics such as an adipose head crest and an anal gland in the first two rays of the anal fin (Fishelson, 1963; Patzner,


Seiwald, Adlgasser, & Kaurin, 1986). For the abovedescribed population in southern Portugal where nest sites are scarce, females court bourgeois males intensively and males assume a passive posture during courtship and often reject females (Almada et al., 1995). The female courtship display is elaborate and markedly different from male displays, and consists of beating the pectoral fins and opening and closing the mouth in synchrony while simultaneously displaying a conspicuous nuptial coloration (Fishelson, 1963; Patzner et al., 1986). Small males, morphologically similar to females, reproduce by mimicking the complex female reproductive displays and nuptial coloration in order to approach large nesting males and achieve parasitic fertilizations of eggs (Gonçalves et al., 1996) (Figure 7.4). Bourgeois males attack and court matched-for-size parasitic males and females with the same probability, suggesting that they are deceived by these parasitic males (Gonçalves, Matos, Fagundes, & Oliveira, 2005). A markrecapture study revealed that the parasitic reproductive tactic is only adopted during the males’ first breeding season, with parasitic males switching into bourgeois males in their second breeding season onwards (Fagundes, Saraiva, Gonçalves, & Oliveira, unpublished data). Thus, the same animal experiences a dramatic change in its sexual behavior, displaying female-like sexual behaviors early in its development and typical male behaviors later on. The above overview of the patterns of sexual behavior in fishes highlights how sexual displays may differ markedly across species, populations, different individuals of the same population, or even within the same animal

(a)

(b)

(c) FIGURE 7.4 Male alternative reproductive tactics in the peacock blenny Salaria pavo. Parasitic males (b) reproduce by mimicking the female (c) morphology and behavior in order to approach the nests of larger nesting males (a) and fertilize eggs. See color plate section.

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throughout its lifetime. This diversity offers an obvious difficulty when trying to derive general physiological mechanisms for the regulation of sexual behaviors but also provides the opportunity to understand the evolution and organization of neuroendocrine systems underlying the expression of sexual displays. Comparative approaches across different systems may prove a useful basis for understanding both the commonalities and variability of the mechanisms underlying male and female sexual displays. However, although a considerable amount of descriptive data on the ethology of sexual behaviors exists for many fish species, neuroendocrine studies focusing on sexual displays are comparatively scarce and research effort has been put into a reduced number of species. This limits for now the usefulness of comparative approaches, and data for more species are required. With this limitation in mind, an overview of what is known about the neuroendocrine regulation of male and female sexual displays in teleost fishes is presented.

3. ENDOCRINE MECHANISMS REGULATING SEXUAL BEHAVIOR Hormones were classically seen as causal agents of behavior where the presence of a particular hormone was necessary for the expression of a given behavior. However, a deeper understanding of the molecular mechanisms through which hormones influence behavior has changed this view into considering hormones as modulators rather than causal agents of behavior (see Oliveira, 2004 for an historical background). By acting on sensory and motor systems and on the central neural circuitry underlying the expression of specific behavioral patterns, hormones, like other modulators, increase or decrease the probability of pattern occurrence. This view helps us to understand apparently paradoxical observations, such as that the expression of specific displays promoted by specific hormones may not be completely abolished after suppressing hormonal action. In teleosts, similarly to other vertebrates, the neuroendocrine system is hierarchically organized, with the hypothalamus controlling the activity of the adenohypophysis, which in turn regulates peripheral endocrine glands. The principal difference between the teleost endocrine system and that of other vertebrates is the lack of a hypothalamice hypophysial portal vascular system to transport the hypothalamic releasing factors to the adenohypophysis. Instead, the adenohypophysis is controlled by direct innervation by hypothalamic neurosecretory neurons (Peter, 1990). For reasons outlined below, studies on the endocrine regulation of fish sexual behavior have focused on hormones of the hypothalamicepituitaryegonadal (HPG) axis, in particular of gonadal steroids, and not much is known about the role

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played by hormones of other axes (e.g., growth hormone, corticosteroids). In some studies sexual behavior has been shown to be independent of gonadal steroids and these observations have promoted the search for other agents potentially involved in the regulation of fish sexual displays. As an example, in the sex-changing bluehead wrasse (Thalassoma bifasciatum) both intact and ovariectomized females responded to the removal of territorial males by occupying their territory and rapidly switching into displaying the full suite of male courtship and spawning behaviors, demonstrating that the action of gonadal steroids was not necessary for the expression of typical male displays (Godwin, Crews, & Warner, 1996; see also Chapter 8, this volume). To account for these observations, the role played by some neuropeptides in the control of fish sexual behavior also has been investigated. An emphasis has been placed on the effects of the gonadotropin-releasing hormone (GnRH) and of the two peptides released by the neurohypophysis, arginine vasotocin (AVT) and isotocin (IST), the fish homologs of mammalian arginine vasopressin (AVP) and oxytocin (OXY), respectively. Our overview on the endocrine mechanisms of fish sexual behavior will thus be focused on this set of neuroendocrine agents.

3.1. Gonadal Steroids Sex steroids have long been recognized as key hormones regulating sexual differentiation, physiological aspects of reproduction, and the development of primary and secondary sexual characteristics (Nelson, 2005). This recognition has established gonadal steroids as prime candidates for the regulation of sexual behavior. The traditional view of the action of sex steroids on behavior was that steroids produced in the gonads exert their effects by acting via nuclear receptors on sensorial, central, or motor output neural systems (Nelson, 2005). However, data suggesting some important exceptions to this general pattern of steroidal action have been accumulating. First, steroids may act via nonclassic membrane receptors (Moore & Evans, 1999). The existence of these receptors was suspected for a long time as steroids were shown to induce physiological and behavioral changes too rapidly to be compatible with their classical genomic action via nuclear receptors (Klein & Henk, 1964; Spach & Streeten, 1964). However, it was not until 2003 that the first membrane steroid receptor, a progestin receptor, was conclusively isolated and cloned in a teleost, the spotted sea trout (Cynoscion nebulosus) (Zhu, Rice, Pang, Pace, & Thomas, 2003). Data on the rapid effects of steroid hormones have been rapidly accumulating over the past years, and a few studies report fast effects of steroid hormones on fish sexual displays (see Section 3.1.1). Second, central mechanisms may regulate the action of

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steroids produced in the periphery. For example, many behavioral actions of T depend on its conversion to estradiol (E2) by the P450 enzyme aromatase (P450aro ¼ CYP19) (e.g., Naftolin et al., 1975). In birds, P450aro has been suggested to be stored in an inactive form in presynaptic boutons and its activity to be quickly regulated by changes in intracellular Ca2þ levels triggered by neurotransmission (Balthazart & Ball, 2006). In a recent study, the rapid modulation of T and E2 concentrations in the brain by social interactions was demonstrated in zebra finches using a microdialysis technique (Remage-Healey, Maidment, & Schlinger, 2008), corroborating this hypothesis. Neural regulation of P450aro activity is thus a possible mechanism to rapidly control local estrogen synthesis from androgens and estrogenic effects in the central nervous system (CNS) (for details see Balthazart & Ball, 2006). Finally, de-novo synthesis of steroids from cholesterol in the brain has been suggested for a few species. For instance, all enzymes necessary for the synthesis of adrenal steroids from cholesterol have been identified in the CNS of mammals (Compagnone & Mellon, 2000). It is thus possible that the regulation of brain circuits underlying the expression of sexual behaviors by steroids is partially independent from their peripheral secretion. Nevertheless, steroids produced in the gonads are still logical candidates for synchronizing gonadal development and gamete maturation with the expression of sexual behavior, and there is ample evidence for their effect on sexual displays. The role of gonadal steroids in fish behavior has been investigated using several complementary approaches. One is to correlate plasma steroid levels with the frequency of behavioral displays (which assumes that the gonads are the main source of the steroids measured in the plasma). Correlational data provide only limited evidence of the role played by specific hormones in specific behavioral displays. For example, in most species dominant males have higher plasma androgen levels and perform aggressive displays more frequently than subordinate males. However, dominant and subordinate males also differ in other aspects (relative testicular development, secondary sexual characteristics, feedback from the social environment) and it is difficult to pinpoint what the functional role of the androgen difference is, based only on correlational data. Since the pioneering experiments of Arnold Berthold (1849), who, by castrating, reimplanting, and transplanting testes in cockerels, demonstrated an effect of a ‘secretory blood-borne product’ in the development of male sexual behavior and morphological traits, castrationehormone replacement experiments have become a standard procedure for testing the effects of gonadal hormones. However, even with this experimental approach it may be difficult to disentangle the effects of hormonal manipulations in the several components of sexual behavior. For instance, a reduction in copulation frequency due to castration and its


restoration by hormone replacement may result from a direct effect of that hormone on copulation behavior or from an effect of the hormone on other aspects of sexual behavior (e.g., nuptial coloration, courtship displays) that indirectly influence copulation success. Direct evidence for the role of a specific hormone in a specific behavior implies demonstrating the site of hormonal action and the mechanism through which the hormone modifies the display. This is only possible using a combination of techniques such as electrophysiological and molecular methods, some of which are technically demanding. For this reason, a detailed knowledge of the way in which specific hormones influence specific sexual behaviors in fishes has been gathered for only a very few well-studied models. With these limitations in mind, a brief overview of the role played by gonadal steroids in male and female sexual behavior is presented below.

3.1.1. Gonadal steroids and male sexual behavior The principal sex steroids detected in the plasma of male teleosts are T, 11b-hydroxytestosterone (OHT), and the nonaromatizable 11-ketotestosterone (11-KT) (Borg, 1994). 11-ketotestosterone is usually the most abundant sex steroid in male plasma and is thought to be the most relevant androgen in male teleosts, whereas T is the major circulating androgen in females (Borg, 1994; Lokman et al., 2002). Similarly, in species with polymorphic male ARTs, bourgeois males have higher plasma levels of 11KT, but not of T, than parasitic males (Oliveira, 2006). Besides playing a role in the regulation of spermatogenesis (e.g., Miura, Yamauchi, Takahashi, & Nagahama, 1991), gonadal differentiation (e.g., Strussmann & Nakamura, 2002), and the development of male secondary sex characteristics (e.g., Mayer, Borg, & Schulz, 1990), gonadal androgens also have been implicated in the activation of male sexual behavior in fishes (for reviews see Liley & Stacey, 1983; Borg, 1994; Oliveira & Gonçalves, 2008). Aggressive behavior associated with reproductive territory acquisition may be facilitated by androgens (e.g., Cardwell & Liley, 1991). However, the available data suggest that the effect of androgen manipulation on male aggression in fishes is largely variable. For example, in sixteen studies where males were castrated, the frequency of aggressive displays decreased in six, remained unchanged in nine, and increased in one study (Table 7.1). Nevertheless, a meta-analysis has confirmed a positive effect of exogenous androgen administration on male aggressiveness in teleosts, and this effect was larger for territorial maleemale aggression than for nest defense aggression (Hirschenhauser & Oliveira, 2006). The central effects of androgens in aggressive behavior are unlikely to depend on T aromatization. Peripheral administration of E2

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TABLE 7.1 Effects of castration on the frequency of aggressive, nest-building, and sexual behaviors in fishes Species

Reference

Aggressive behavior No effect Betta splendens (breeding males)a

Weiss and Coughlin (1979)

Macropodus opercularis (breeding males)a

Villars and Davis (1977)

Trichogaster trichopterus (breeding males)

Johns and Liley (1970)

Gasterosteus aculeatus (pre-nesting males; short-days)

Hoar (1962)

Gasterosteus aculeatus (pre-nesting males)

Baggerman (1966); Wootton (1970)

Lepomis megalotis (territorial males)

Smith (1969)

Lepomis gibbosus (territorial males)

Smith (1969)

Salaria pavo (parasitic males)

Gonçalves et al. (2007)

Increased Gasterosteus aculeatus (pre-nesting males; long-days)

Hoar (1962)

Decreased Gasterosteus aculeatus (nesting males)

Baggerman (1966); Wootton (1970)

Astatotilapia burtoni (males)

Francis et al. (1992)

Pseudocrenilabrus multicolor (males)

Reinboth and Roxner (1970)

Xiphophorus maculatus (males)

Chizinsky (1968)

Bathygobius soporator (breeding males)

Tavolga (1955)

Nest building No effect Sarotherodon melanotheron melanotheron (breeding males)

Aronson (1951)

Sarotherodon melanotheron heudelotii (breeding males)

Heinrich (1967)

Oreochromis upembae (breeding males)

Heinrich (1967)

Gasterosteus aculeatus (breeding males)

Ikeda (1933)

Macropodus opercularis (breeding males)

Villars and Davis (1977)

Decreased Lepomis gibbosus and L. megalotis (breeding males)

Smith (1969)

Pseudocrenilabrus multicolor multicolor (breeding males)

Reinboth and Rixner (1970)


Baggerman (1957); Hoar (1962); Wai and Hoar (1963); Baggerman (1966); Borg (1987)



Sexual behavior No effect Hemichromis bimaculatus (breeding males)

Noble and Kumpf (1936)


Baggerman (1968); Pa´ll et al. (2002)

Bathygobius soporator (breeding males)

Tavolga (1955)

Thalassoma bifasciatum (breeding males)

Semsar and Godwin (2003)

Decreased Pseudocrenilabrus multicolor multicolor (breeding males)

Reinboth and Rixner (1970)


Baggerman (1957); Hoar (1962) (Continued)

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TABLE 7.1 Effects of castration on the frequency of aggressive, nest-building, and sexual behaviors in fishesdcont’d Species

Reference

Morone americana (breeding males)

Salek (2001)



Oncorhynchus mykiss (breeding males)

Mayer et al. (1994)

a

Incomplete gonadectomy or evidence for rapid testicular regeneration.

has an inhibitory effect on male aggression in fishes (Bell, 2001; Clotfelter & Rodriguez, 2006; Gonçalves, Alpedrinha, Teles, & Oliveira, 2007). In a sex-changing goby, brain P450aro activity decreases with aggression, suggesting that a decrease in E2 synthesis (or a higher availability of T) facilitates aggressive behavior (Black, Balthazart, Baillien, & Grober, 2005). Both T and 11-KT promote aggression in teleosts (Oliveira & Gonçalves, 2008), although in a recent study pre-fight levels of T but not of 11-KT correlated positively with aggressive displays in the internally selffertilizing hermaphroditic killifish, Kryptolebias marmoratus (Earley & Hsu, 2008). The role of gonadal steroids in nest-building behavior has been investigated in only a few species but in greatest detail in G. aculeatus. The results from twelve castration studies in several species range from no effect (five studies) to a suppression of nest-building behavior (seven studies) (Table 7.1). Both T and 11-KT administration to castrates restores the behavior (Oliveira & Gonçalves, 2008), and in sticklebacks 11-ketoandrostenedione (11-KA) seems to be more effective than T in restoring nest-building behavior (Borg, 1987). In G. aculeatus, males exposed to E2 start building nests later, although E2 does not affect the percentage of males that build nests (Wibe, Rosenqvist, & Jenssen, 2002). Also in this species, the males’ exposure to the antiandrogen flutamide decreases nest-building behavior at low concentrations of the substance and completely abolishes the behavior at higher levels, although this may be a consequence of lower spiggin production at these concentrations (Sebire, Allen, Bersuder, & Katsiadaki, 2008). Similarly, exposure to fenitrothion, an organophosphorus pesticide structurally similar to flutamide, reduces spiggin production and nest-building behavior in this species (Sebire et al., 2009). Taken together, the available data suggest that the central action of androgens secreted by the testis promotes nest-building behavior in males. Hormonal manipulations also induce largely variable effects in male courtship displays. In eleven studies in which males were castrated, the frequency of male sexual displays was maintained in five and reduced in six cases (Table 7.1). In five out of these six cases, some castrated

males were also administered with androgens. Testosterone (two studies), methyltestosterone (MT) (one study), and 11-KT (one study) at least partially recovered male sexual displays, and in one case 11-KA failed to do so. In intact males, T administration failed to promote male sexual behavior in four different species whereas 11-KT or 11-KA promoted male sexual displays in two out of four species. Blocking androgen action with cyproterone acetate, flutamide, or vinclozolin reduced male sexual displays in four out of five cases (Oliveira & Gonçalves, 2008). Hormonal manipulations in females and immature males have also produced variable results. As an example, female goldfish treated with 11-KT (but not with T) displayed male-like sexual behaviors in response to a stimulus female (Stacey & Kobayashi, 1996), while in G. aculeatus 11-KA (Borg & Mayer, 1995) or MT (Wai & Hoar, 1963) administration to females failed to induce male-like behaviors. Further, the role of central aromatization of T into E2 in the regulation of male sexual displays, well established in mammals and birds (reviewed in Baum, 2003; Ball & Balthazart, 2004), is not clear in fishes. In a study in guppies (P. reticulata) two of three male sexual displays were reduced by P450aro inhibition (Hallgren, Linderoth, & Olsen, 2006), suggesting that, similar to other vertebrates, T aromatization to E2 facilitates some aspects of male sexual behavior. On the other hand, for five species in which exogenous E2 was administered to males, sexual displays were reduced in four cases and remained unchanged in one case (Oliveira & Gonçalves, 2008). Moreover, in weak electric fish, androgens masculinize the EOD signal in females, castrated males, juvenile males, or males in nonreproductive conditions (e.g., Bass & Hopkins, 1983; Meyer, 1983; Bass & Hopkins, 1985; Bass & Volman, 1987; Dunlap & Zakon, 1998; Herfeld & Moller, 1998; Few & Zakon, 2001) whereas estrogens feminize the signal (Dunlap, McAnelly, & Zakon, 1997), probably via transcriptional regulation of ionic membrane channels in electrocytes, the cells of the electric organ (e.g., Liu, Wu, & Zakon, 2008). Thus, the role of central T aromatization in the regulation of male sexual behavior in fishes is for now unclear and more experiments are necessary to assess its relevance.

Chapter | 7


A role for progestins in male courtship displays also has been suggested. Plasma levels of the progestin 17,20b-dihydroxy-4-pregen-3-one (17,20bP) correlate with male courtship displays in several species (e.g., Oncorhynchus mykiss (Liley, Breton, Fostier, & Tan, 1986), Salmo salar (Mayer et al., 1990b), Salmo trutta (Olseń, Ja¨rvi, Mayer, Petersson, & Kroon, 1998), and Chromis dispilus (Barnett & Pankhurst, 1994)). Further, in species with male ART, differences in circulating levels of the progestins 17,20bP, 17,20b,21-trihydroxy-4-pregen-3-one (17,20b,21P), and 17,20a-dihydroxy-4-pregen-3-one (17,20aP) occur (for a review see Oliveira, 2006). However, because progestins play a role in reproductive functions, namely in spermatogenesis and spermiation (e.g.,Miura, Higuchi, Ozaki, Ohta, & Miura, 2006), it is difficult to assess whether these differences are a consequence of the progestins’ actions in the testis and/or whether they reflect the central action of progestins in the modulation of male sexual displays. Evidence for a central regulation of behavior by progestins comes from a study in the rainbow trout (O. mykiss), where the administration of 17,20bP, but not of 11-KA, to castrated males restored male sexual behavior (Mayer, Liley, & Borg, 1994). Additionally, a pheromonal role for progestins has been suggested. In females, 17,20bP plasma levels peak prior to ovulation and are released into the water. Recently, putative progestin receptor genes with potential to respond to steroid pheromones have been identified in the goldfish olfactory epithelium transcriptome (Kolmakov, Kube, Reinhardt, & Cana´rio, 2008), and 17,20bP has been shown to act as a potent stimulator of the males’ olfactory system (e.g., Sorensen, Scott, Stacey, & Bowdin, 1995). In contrast, exposure of male goldfish to 17,20bP induced an increase in gonadotropin (GTH) levels and milt production but had only minor effects on behavior (Sorensen, Stacey, & Chamberlain, 1989), and thus the putative pheromonal role of 17,20bP needs to be further investigated (see Chapter 6, this volume). Rapid effects of steroid manipulation on behavior, consistent with a nongenomic action, have also been documented in fishes. In both the Gulf toadfish (Opsanus beta) and the closely related plainfin midshipman (Porichthys notatus), nesting males (type I) produce vocalizations to attract females into their nests, and in P. notatus parasitic female-like males (type II) that do not emit mating calls also have been described. In these species, it is possible to study the pattern of ‘fictive vocalizations’ in electrophysiological preparations. Electrical microstimulation in forebrain/midbrain vocal nuclei induces a rhythmic motor volley that can be intracranially recorded from occipital nerve roots descending into the sonic muscles. As the temporal pattern of the vocal motor volley determines the properties of the natural calls, the vocal motor volley is designated as a ‘fictive vocalization’ (Bass,

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2008; see below for details). Intramuscular injections of 11-KT increased, within minutes, the duration of fictive vocalizations in both species and this effect was restricted to type I males, the morphotype with higher plasma 11-KT levels (Brantley, Wingfield, & Bass, 1993; Remage-Healey & Bass, 2004; 2006a; 2007). Fictive vocalizations in females of both species and in type II males of P. notatus were rapidly responsive to T instead of 11-KT, and T is the main circulating androgen in these morphs (Remage-Healey & Bass, 2007). This is consistent with a field experiment in toadfish where 11-KT administered to males (using food containing 11-KT crystals) increased their calling behavior within minutes (Remage-Healey & Bass, 2006a; for reviews see Remage-Healey & Bass, 2006b; Bass & Remage-Healey, 2008).

3.1.2. Gonadal steroids and female sexual behavior In females, both E2 and T are major circulating steroids, while 11-KT, the main androgenic steroid in males, is usually undetectable or present only in low concentrations (Borg, 1994 but see Lokman et al., 2002; Desjardins, Hazelden, Van der Kraak, & Balshine, 2006). Stacey (1981) has proposed that prostaglandins (PGs) produced by mature oocytes in external fertilizing species may signal a ready-to-spawn state and thus promote female sexual behaviors. In internal fertilizers, sexual behavior and fertilization are temporally dissociated and estrogens produced during follicular development may induce sexual behaviors in anticipation of ovulation. Although studies on the endocrine regulation of female sexual behavior in fishes are scarce, effects of gonadal steroids on various aspects of female sexual displays in both internal and external fertilizers have been reported. In the cichlid Neolamprologus pulcher, the male and the female jointly defend their breeding territory. Females are more aggressive than males and also have higher plasma T levels than their mates, suggesting a role for T in female aggression (Desjardins et al., 2008). Accordingly, in the blue acara (Aequidens pulcher), T increases and E2 decreases the frequency of female aggressive behaviors (Munro & Pitcher, 1985). In the fighting fish (B. splendens), daily injections of T to females for nine weeks progressively increased aggressiveness towards males but decreased aggressiveness towards females (Badura & Friedman, 1988). Androgens also induce nest-building behavior in females in species where only males typically build nests. For instance, in two cichlids (Astotilapia burtoni and Pseudocrenilabrus multicolor multicolor), T induces nest-building behavior in females (Reinboth & Rixner, 1970; Wapler-Leong & Reinboth, 1974). However, the role of gonadal steroids in this aspect of female sexual behavior in species where females naturally exhibit nest

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building has not been investigated. These results suggest that in females, androgens, particularly T, promote aggressive and nest-building behaviors. Females take a passive role during courtship and lack stereotyped courtship displays in many teleost fishes. As a consequence, the neuroendocrine control of female courtship displays has been poorly investigated but sex-rolereversed species offer the possibility to overcome this difficulty. In the above-described sex-role-reversed population of the peacock blenny in southern Portugal, females take the initiative in courtship and exhibit a stereotyped courtship display and a typical nuptial coloration when approaching a male’s nest (Almada et al., 1995). The principal circulating sex steroid in females is E2 and both T and 11-KT levels are significantly lower than in nesting males (Gonçalves, Teles, Alpedrinha, & Oliveira, 2008). To test the roles of gonadal hormones in female sexual behavior, females were either sham-operated, ovariectomized and implanted with vehicle, or ovariectomized and implanted with E2, and their courtship behavior towards a nesting male was tested for one hour every two days for fourteen days. Ovariectomy significantly reduced the frequency of the female courtship displays although 75% of ovariectomized vehicle-implanted females still courted a nesting male at least once after 14 days in comparison with approximately 90% of sham-operated females. Estrogen partially restored female sexual behavior as the frequency of courtship displays and the time spent in nuptial coloration was intermediate in E2-treated females when compared with shamoperated and vehicle-implanted females (Gonçalves, Teles, Costa, and Oliveira, unpublished data). These results suggest that E2 produced in the ovaries promotes female sexual behavior in S. pavo but also show that the behavior may persist after 14 days without gonadal steroid production. In this species, female-like sneaker males mimic the females’ displays, and the effects of steroids in the sneakers’ behavior also have been tested. Estradiol implants to castrated sneakers failed to promote female-like displays (Gonçalves et al., 2007). Interestingly, castration increased female-like displays while T or 11-KT implants decreased the frequency of these displays, suggesting an inhibitory role for androgens in the sneakers’ female-mimicking behavior (Oliveira, Carneiro, Gonçalves, Cana´rio, & Grober, 2001; Gonçalves et al., 2007) (Figure 7.5). These results show that similar courtship displays in females and sneakers may be under the regulation of different endocrine mechanisms. The results presented above for males and females stress the enormous variability in the effects of endocrine manipulation on appetitive aspects of sexual behavior in teleost fishes. It seems likely that part of this variability reflects interspecific variation in the mechanisms of hormonal regulation of sexual behavior, but some is certainly derived from methodological differences between studies. For example, an inverted U-shaped function has


FIGURE 7.5 Frequency of female-like displays directed by parasitic males towards a nesting male. Parasitic males were either castrated, sham operated, castrated and implanted with testosterone, or castrated and implanted with estradiol. Adapted from Gonçalves, Alpedrinha, Teles, and Oliveira (2007).

been reported for the effects of cortisol and AVT on fish behavior, with maximal stimulatory effects occurring at intermediate dosages of AVT (Santangelo & Bass, 2006) or cortisol (Bass, 2008; see also Volkoff & Peter, 1999). In most studies investigating the behavioral effects of hormonal manipulations, dose-response curves are not considered and only a single dosage is used. Thus, hormonal concentrations, time between hormonal manipulation and behavioral observations, or methods of hormone delivery are just a few of the many technical aspects that vary between studies and that make their comparison difficult. The field would certainly benefit from standardizing some of these procedures.

3.2. Neuropeptides 3.2.1. Gonadotropin-releasing hormone (GnRH) Gonadotropin-releasing hormone is a highly conserved decapeptide that appears in three forms in vertebrates: GnRH1, GnRH2, and GnRH3. A gene duplication event early in the teleost lineage gave rise to the third isoform, GnRH3 (White, Kasten, Bond, Adelman, & Fernald, 1995; Parhar, 1997). The three isoforms have different distributions in the teleost brain and perform different functions (for reviews see Hofmann, 2006; Chen & Fernald, 2008; see also Chapter 2, this volume). GnRH1 neurons occur mainly in the preoptic area (POA)/anterior hypothalamus and project to the adenohypophysis, regulating the activity of the gonadotrope cells that synthesize follicle-stimulating hormone (FSH) and luteinizing hormone (LH). Release of FSH and LH into the bloodstream stimulates normal gonadal development and gonadal steroid production in both males and females, and as we have seen gonadal

Chapter | 7


steroids are relevant modulators of sexual behavior. Alternatively, GnRH2 and GnRH3 may directly modulate sexual behavior by acting on central mechanisms. GnRH2 is found throughout the brain, especially in the midbrain system, and is thought to play a neuromodulatory role. This isoform stimulates female sexual behavior in mammals (Barnett, Bunnell, Millar, & Abbott, 2006) and fishes (Volkoff & Peter, 1999). In the goldfish brain, mRNA expression levels of GnRH2 correlated with female spawning behavior (Canosa, Stacey, & Peter, 2008) and intracerebroventricular injections of low doses of GnRH2 and GnRH3 stimulated female spawning behavior, while higher dosages inhibited the behavior (Volkoff & Peter, 1999). GnRH3 neurons are found in the terminal nerve system with projections throughout the brain and this peptide is also thought to act as a neuromodulator (Oka & Matsushima, 1993). Lesioning of the terminal nerve impairs nest-building behavior in male dwarf gouramis (Colisa lalia) (Yamamoto, Oka, & Kawashima, 1997). In male tilapia (Oreochromis niloticus) immunoneutralization of GnRH3 (but not of GnRH1 or GnRH2) has the same suppressive effect on male nestbuilding behavior and also reduces aggressive displays (Ogawa et al., 2006). It seems possible that the effects of gonadal steroids on fish sexual behavior mentioned above are partly the result of an interaction with GnRH neurons, and several studies report a modulation of GnRH by sex steroids (e.g., Amano et al., 1994; Parhar, Tosaki, Sakuma, & Kobayashi, 2001; Levavi-Sivan, Biran, & Fireman, 2006; Vetillard, Ferriere, Jego, & Bailhache, 2006).

3.2.2. Arginine-vasotocin (AVT) Besides its well-established role in osmoregulation, the nonapeptide AVT has been implicated in the regulation of social and sexual behavior in vertebrates (for review see Foran & Bass, 1999; Goodson & Bass, 2001; Balment, Lu, Weybourne, & Warne, 2006). In teleost fishes, AVT neurons occur in the POA and project to the neurohypophysis and to many other brain regions. Within the POA, three subpopulations of AVT neurons can be identified: parvocellular, magnocellular, and gigantocellular, and different roles for these populations have been suggested (e.g., Greenwood, Wark, Fernald, & Hofmann, 2008). Arginine vasotocin has been implicated in male aggression in fishes, though with variable effects. For example, in the bluehead wrasse, AVT administration to territorial terminal phase (TP) males decreased aggression towards initial phase (IP) males while AVT administration to nonterritorial terminal phase males had the opposite effect (Semsar, Kandel, & Godwin, 2001) (see Chapter 8, this volume, for discussion of TP and IP in sex-changing fishes). In another field study, AVT injections to territorial male beaugregory damselfish (Stegastes leucostictus) promoted aggressive behavior towards intruders at medium

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dosages but had no effect at low and high dosages (Santangelo & Bass, 2006). In several other species, exogenous AVT administration decreased aggressive behavior. Arginine vasotocin reduced the production of an EOD signal used in agonistic contexts in the weakly electric fish Apteronotus leptorhynchus (Bastian, Schniederjan, & Nguyenkim, 2001), and decreased aggressive behavior in the Amargosa pupfish (Cyprinodon nevadensis amargosae) (Lema & Nevitt, 2004) and in juvenile rainbow trout (O. mykiss) (Backstro¨m & Winberg, 2009). In the plainfin midshipman, type I territorial males use grunt vocalizations as aggressive acoustic displays (Brantley & Bass, 1994) and AVT inhibited induced fictive grunt vocalizations in electrophysiological preparations (Goodson & Bass, 2000a; 2000b) (Figure 7.6). In A. burtoni, AVT mRNA expression in the gigantocellular layer was higher in territorial than in nonterritorial males, while in the parvocellular layer an opposite pattern was found (Greenwood et al., 2008). The authors of the study suggest that this difference may relate to a role of the gigantocellular AVT neurons in the promotion of dominance traits (aggression, courtship, and/or upregulation of reproductive physiology) in contrast to the activation of subordinate behavior or of the stress axis by parvocellular AVT neurons. If this is the case, exogenous administration of AVT by implants or injections may have unexpected effects as the targets of both gigantocellular and parvocellular neurons are activated. This is corroborated by the between- and within-species variable effects of AVT manipulation on the regulation of aggressive behavior in teleosts and in other vertebrates (Goodson & Bass, 2001). Arginine vasotocin also has been implicated in the regulation of courtship behavior. Females of the bluehead wrasse may undergo sex change when a territorial TP male is removed (Warner & Swearer, 1991) and, as mentioned above, the female-to-male behavioral switch is fast and independent of the gonads (Godwin et al., 1996). Arginine vasotocin mRNA expression increases in the brain of sex-changing females (Godwin, Sawby, Warner, Crews, & Grober, 2000) and is higher even in ovariectomized dominant sex-changing females when compared with subordinate females (Semsar & Godwin, 2003). Arginine vasotocin administration to both territorial and nonterritorial TP males promoted male courtship behavior, further supporting an activational role of AVT in male sexual displays in this species (Semsar et al., 2001). However, this activational role seems to be context-dependent, as AVT administration to IP males or females fails to induce male-like behaviors under social conditions that inhibit sex change (i.e., in the presence of a territorial TP male) (Semsar & Godwin, 2004; see also Chapter 8, this volume). Arginine vasotocin facilitates male sexual displays in other fish species. In the weakly electric fish A. leptorhynchus, AVT increased the production of electrical

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FIGURE 7.6 (a) Schematic representation of the prepacemaker nucleus and electric organ in a gymnotiform. Redrawn from McAnelly and Zakon (2000). (b) The neural circuitry regulating electric organ discharge (EOD) in gymnotiforms. The pacemaker nucleus (PMN) sets the EOD frequency. The firing rate of the PMN is in turn modulated by the thalamic prepacemaker nucleus (PPn) and by the sublemniscal prepacemaker nucleus (SPPn). Redrawn from Zakon, Oestreich, Tallarovic, and Triefenbach (2002). (c) The type I male midshipman produces vocalizations by vibrating the swim bladder lateral walls through a pair of sonic muscles. Reproduced from Bass and Zakon (2005). (d) The vocal motor network includes vocaleacoustic integration centers (VACs) at the forebrain (f), midbrain (m), and hindbrain (h) level that input, via a ventral medullary nucleus, into the hindbrainespinal sonic motor nucleus, which innervates the sonic muscles (for details and other references see Bass and Zakon, 2005).

signals used in female attraction (Bastian et al., 2001), and in male white perch (Morone americana) intracerebroventricular AVT administration promoted male sexual displays (Salek, Sullivan, & Godwin, 2002). In the grass puffer (Takifugu niphobles), brain AVT mRNA expression increases in prespawning females, suggesting a relationship with female sexual behavior (Motohashi, Hamabata, & Ando, 2006). In the above-mentioned sexrole-reversed population of the peacock blenny (S. pavo), AVT mRNA was higher in the courting morphs (females and female-like sneaker males) than in nest-holders, suggesting that AVT is correlated with the expression of female and female-like courtship displays (Grober, George, Watkins, Carneiro, & Oliveira, 2002). Accordingly, exogenous AVT administration to females and sneakers promoted female courtship displays, whereas AVT administration had no effect on the sexual displays by nest-holders (Carneiro, Oliveira, Cana´rio, & Grober, 2003). Taken together, these results suggest that AVT generally promotes courtship behavior in teleost fishes, but future studies are necessary to clarify how the different subpopulations of AVT neurons regulate male and female sexual behavior. It is also possible that the modulation of courtship behavior by the AVT system is under the regulation of GnRH as, in O. mykiss, in-vitro GnRH administration to POA-AVT neurons stimulated their electrical activity (Saito, Hasegawa, & Urano, 2003).

3.2.3. Isotocin (IST) Oxytocin, the mammalian homolog of IST, has been implicated in the activation of female sexual behavior and in the inhibition of female aggression in rats (Pedersen &

Boccia, 2002). The effects of IST on fish sexual behavior, however, have been investigated in only a few studies. In electrophysiological studies of the plainfin midshipman CNS, IST inhibited fictive grunt vocalizations (used in an agonistic context) in type II parasitic males and females but not in type I territorial males (Goodson & Bass, 2000a). In the beaugregory damselfish (S. leucosticus), intramuscular injections of IST had no effect on territorial male aggressive behavior (Santangelo & Bass, 2006). In the goldfish, intraperitoneal injections of IST increases plasma E2 levels (Mennigen et al., 2008), and E2 inhibits aggressive behavior in fish (e.g., Gonçalves et al., 2007). The results thus suggest that IST reduces aggressive behavior in fishes, and there is the possibility that these effects are partially mediated by steroids. In females of the grass puffer, brain IST mRNA expression did not change during the breeding period (Motohashi et al., 2006), although brain IST levels measured with high performance liquid chromatography (HPLC) in female (but not male) three-spined stickleback were maximal during the reproductive season (Gozdowska, Kleszczynska, Sokolowska, & Kulczykowska, 2006). In spite of these correlational data, the role of IST in fish courtship displays is for now unclear as experimental studies are missing. Neurohypophysial hormones are also thought to play a role in consummatory aspects of fish sexual behavior. The spawning reflex has been induced by the administration of pituitary extracts to hypophysectomized male killifish (Fundulus heteroclitus) (Pickford, 1952) and this result was later confirmed for both males and females of the same species (Wilhelmi, Pickford, & Sawyer, 1955). Similar results were afterward obtained for female medaka

Chapter | 7


(Oryzias latipes) (Egami, 1959), female bitterling (Rhodeus sericeus) (Egami & Ishii, 1962), and male and female flagfish (Jordanella floridae) (Crawford, 1975). In the killifish, AVT was more effective than IST in eliciting the spawning reflex (Pickford & Strecker, 1977). In several other species, however, neurohypophysial hormones failed to induce the spawning reflex (G. aculeatus (Lam and Nagahama, personal communication in Liley & Stacey, 1983), C. auratus (Pickford in Macey, Pickford, & Peter, 1974), Misgurnus fossilis, and S. salar (Egami & Ishii, 1962)). In male seahorses (Hippocampus hippocampus), both OXY and IST failed to induce spawning behavior, although these substances did induce parturition reflexes (Fiedler, 1970). Interestingly, in Clarias batrachus intraovarian pressure induces the hypertrophy of POA neurosecretory neurons (Subhedar, Krishna, & Deshmukh, 1987) and pharmacologically blocking putative mechanosensory channels in the ovary inhibits the effect in POA cells (Subhedar, Deshmukh, Jain, Khan, & Krishna, 1996; see also Deshmukh & Subhedar, 1993). This could be a potential mechanism for synchronizing ovarian development and oocyte maturation with the putative endocrineinduced spawning reflex.

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facilitation of female reproductive behavior by PGF2a has been extended to other species, namely the cichlid Cichlasoma bimaculatum (Cole & Stacey, 1984), the paradise fish (Macropodus opercularis) (Villars, Hale, & Chapnick, 1985), and the Java barb (Barbonymus gonionotus) (Liley & Tan, 1985). In spite of this well-established role of PGF2a in facilitating female sexual behavior, the neural targets of PGF2a responsible for this effect are, for now, unknown. Additionally, PGF2a produced by ovulated females is released into the water and acts as a potent olfactory stimulant for males (Sorensen, Hara, Stacey, & Goetz, 1988; Sorensen et al., 1989). Exposure to PGF2a upregulates GnRH3 mRNA expression in the male brain (ChungDavidson, Rees, Bryan, & Li, 2008) and thus the pheromonal action of PGF2a on male sexual displays may be mediated by a GnRH system. By promoting receptivity and spawning behavior in females and activating sexual responses in males, PGF2a produced in the oviduct of mature females is thought to synchronize male and female reproductive behavior (for reviews see Kobayashi et al., 2002; Stacey, 2003).

3.3. Prostaglandins (PGs)

4. BRAIN CIRCUITS UNDERLYING SEXUAL BEHAVIOR IN FISHES

In mammals, the fatty-acid-derived PGs promote luteolysis, facilitate female sexual behavior (e.g., Buntin & Lisk, 1979), and mediate the perinatal brain masculinization induced by steroids (Amateau & McCarthy, 2004). In fishes, plasma PGs, in particular PGF2a, peak during ovulation (e.g., Goetz & Cetta, 1983). Prostaglandins produced in the mature ovary may serve three main functions in teleosts: to act in a paracrine fashion to stimulate follicular rupture; to enter the bloodstream and act on the nervous system, activating female sexual receptivity and spawning behavior; and to be released into the water as a pheromone in order to stimulate male sexual behavior (for a review see Sorensen & Goetz, 1993; see also Chapter 6, this volume). The role of PGF2a in sexual behavior has been investigated in more detail in the goldfish. Injecting non-ripe female goldfish with mature oocytes (Stacey & Liley, 1974) or with PGF2a (e.g., Stacey & Liley, 1974; Stacey, 1976; 1981; Sorensen & Goetz, 1993; Kobayashi, Sorensen, & Stacey, 2002) stimulates female behavior associated with oviposition. Response to PGF2a administration is similar in ovariectomized, intact, or sham females, suggesting that oviposition behavior is probably induced by PGF2a produced in the reproductive tract during ovulation (Kobayashi & Stacey, 1993). Additionally, sexually mature male goldfish treated with PGF2a display female-like spawning behavior similar to PGF2a-injected females (Stacey, 1976; 1977; Stacey & Kyle, 1983). The

To understand how hormones modulate sexual behaviors it is fundamental to identify the neural substrates upon which hormones act. The techniques available to study the brain areas implicated in the expression of behaviors in fishes include electrophysiological and immunohistochemical techniques, gene expression, directed lesions, and more recently in-vivo imaging and gene manipulation techniques (see Section 5). Since the 1950s, ablation and lesion studies have shown that telencephalic regions are important for the expression of sexual behaviors. Bilateral ablation of the telencephalon does not abolish sexual behaviors but reduces the frequency of their expression (e.g., Kamrin & Aronson, 1954; Aronson & Kaplan, 1968; Davis & Kassel, 1980). Studies in which partial lesions directed to various regions of the telencephalon were conducted suggest that the suppressive effects of bilateral telencephalon ablation on sexual behavior are a consequence of damage to the ventral regions of the telencephalon, including those containing part of the POA (e.g., Davis, Kassel, & Martinez, 1981; Koyama, Satou, Oka, & Ueda, 1984). Lesions to the POA reduce the male spawning reflex in F. heteroclitus (Macey et al., 1974) and lesions to ventral telencephalon regions dorsal and anterior to the anterior commisure (the area ventralis telencephali pars supracommisuralis and the posterior part of the area ventralis telencephali pars ventralis) impair male spawning behavior in the goldfish (Kyle & Peter, 1982; Kyle, Stacey, & Peter, 1982). Electrical

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stimulation of these brain areas, on the other hand, evokes courtship and spawning behaviors in several species (e.g., Demski & Knigge, 1971; Demski, Bauer, & Gerald, 1975; Demski & Hornby, 1982; Satou et al., 1984). In spite of these early studies and of the more recent technical possibilities, detailed knowledge on the neural circuitry underlying the expression of specific sexual behaviors is available for only a few species. The mechanism of EOD signal production of electric fishes and the sound-emitting behavior of sound-producing fishes are probably the bestdescribed models so far (for a comparative review see Bass & Zakon, 2005). The wave-type EOD signal of Gymnotiformes and the vocal signaling behavior of the plainfin midshipman are presented as examples, although other electric (i.e., mormyrids) and acoustic (e.g., toadfish) systems have been well characterized. The electrical signal of Gymnotiformes is produced by electrocyte cells displayed bilaterally in a serial arrangement in the tail. These cells are activated by electromotor neurons, which, in turn, receive input from relay cells adjacent to a small set of ca. 50e100 neurons located in the pacemaker nucleus (Pn) of the medulla oblongata (Figure 7.6). The pacemaker neurons discharge spontaneously and set the EOD timing in a 1 : 1 fashion. The pulse duration of the EOD, on the other hand, is determined by the ion current kinetics of the electrocytes. Within the same animal, consecutive EODs fire at a highly regular rate, and the wave-type EOD signal of Gymnotiformes is probably the most regular pacemaker system known in vertebrates (Moortgat, Keller, Bullock, & Sejnowski, 1998). This continuous firing serves the role of electrosensing, but the properties of the EOD waveform also communicate species, sex, and social status (Wong & Hopkins, 2007). Besides this function, the EOD signal is used in communication by changing its temporal and spectral properties. For example, as part of their courtship displays, males of the genus Brachyhypopomus increase the frequency of the EODs in early phases of the courtship sequence (‘acceleration’) and reduce the amplitude of the EODs in later phases (‘chirp’) (for review see Stoddard, 2002; Stoddard, Zakon, Markham, & McAnelly, 2006). The Pn receives input from two other nuclei: the midbrain sublemniscal prepacemaker nucleus (SPPn) and the thalamic prepacemaker nucleus (PPn) (Figure 7.6). In Eigenmannia, cells from the SPPn mediate smooth decreases in the frequency of the EOD. Within the PPn, two subpopulations of neurons have been identified: a medial population that mediates gradual increases in EOD frequency (PPn-G) and lateral magnocellular neurons (PPn-C) that evoke interruptions in the EOD. Prepacemaker nuclei in turn receive input from the ventral telencephalon and POA (reviewed in Zupanc & Maler, 1997; Zupanc, 2002). In Eigenmannia, electrical stimulation of rostral POA regions, but not of other forebrain regions, evoke interruptions in the EOD discharges,


similar to those naturally observed during courtship, suggesting that the POA may control some aspects of the EOD modulation observed during courtship (Wong, 2000). The wave-type EOD signal is sexually dimorphic, with the EOD pulse being longer in males than in females (e.g., Hopkins, Comfort, Bastian, & Bass, 1990). Additionally, rapid, intermediate, or long-term changes in the properties of the EOD occur and are regulated by different neuroendocrine mechanisms. For example, the rapid production of chirps in A. leptorhynchus is mediated by AVT (Bastian et al., 2001); the intermediate (6e45 minute) changes in EOD properties observed during agonistic interactions in B. pinnicaudatus are mediated by melanocortin peptide hormones such as corticotropin (ACTH) (Markham & Stoddard, 2005); and the long-term changes in the EOD signal (seasonal, ontogenetic) are mediated by sex steroids (e.g., Few & Zakon, 2001; 2007). As androgen administration decreases EOD frequency, set by Pn neurons, and also increases pulse duration, determined by ionic properties of the electrocyte membranes, androgens are predicted to act on both sites. Indeed, electrocytes express androgen receptors (Gustavson, Zakon, & Prins, 1994), small androgen implants in the electric organ increase EOD pulse duration but not frequency (Few & Zakon, 2001), and androgens induce gene expression changes in Kþ ionic channels in electrocytes that underlie the EOD pulse duration increase (Few & Zakon, 2007). Although androgen receptors are not expressed in the Pn (Gustavson et al., 1994), they are located in the upstream SPPn and PPn nuclei (Zakon, 1996), and thus it is possible that androgens masculinize the EOD frequency acting on these nuclei or even in other brain regions afferent to the prepacemaker nuclei. In the plainfin midshipman and in other Batrachoididae, sounds are produced by the contraction of a pair of sonic muscles attached to the swim bladder. Unlike the electrical signal of Gymnotiformes, sounds are only produced in social contexts. The sonic muscles are commanded by ipsilateral motor neurons originating from a bilateral hindbrainespinal cord nucleus. The firing rate of the sonic motor neurons is established by adjacent pacemaker-like neurons that innervate the sonic motor nucleus. Together, the pacemaker and sonic motor neurons form the vocal pattern generator (Figure 7.6). Modulation of the firing rate of pacemaker neurons occurs via more anterior regions. The pacemaker nucleus receives input from a rostro-ventral medullary nucleus and this in turn receives input from vocaleacoustic integration centers (VACs) located in the mid and hindbrain (Figure 7.6). A third forebrain VAC located in the POA/anterior hypothalamus inputs into the midbrain VAC. The synchronous activity of the sonic motor neurons produces compound action potentials that can be easily recorded from the descending occipital nerve roots and that are matched 1 : 1 with the sonic muscle contraction

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and sound pulses. The compound action potentials can be elicited by electric stimulation of forebrain and midbrain VACs (‘fictive calls’) (for details see Bass, 2008). During the breeding season, nesting males produce long-lasting (onehour) advertisement ‘hums’ used in female attraction and short (50e100 ms) defense ‘grunts’ (Bass & McKibben, 2003). The surge in circulating 11-KT observed in nesting males during the breeding season is thought to trigger the ‘hum’ calls (Knapp, Marchaterre, & Bass, 2001). In accordance, intramuscular injections of 11-KT (and also E2 and cortisol) to nesting males caused within five minutes a significant increase in the duration of midbrain-elicited fictive calls (Remage-Healey & Bass, 2004). Contrarily, T is the predominant circulating androgen during the reproductive period in females and sneaker males, and T, but not 11-KT, increases the vocal hindbrain-spinal pattern generator (VPG) output (Remage-Healey & Bass, 2007). Using intact and semi-intact CNS preparations, the hindbrainespinal cord region has been found to be necessary and sufficient for these fast effects of steroids, while midbrain input seems necessary for the continuation (up to 120 minutes) of these effects (Remage-Healey & Bass, 2004). These two examples testify how understanding of the neural circuitry underlying behaviors may help us to draw testable hypotheses related to their endocrine regulation, and how this contributes to an integrated view of the neurophysiological mechanisms of behavior. Nevertheless, the extraordinary research being carried out in electric and acoustic fishes is for the time being not paralleled in other species, and in general our knowledge about the brain areas underlying the expression of sexual behaviors in fishes and their neuroendocrine regulation is still poor.

5. PROSPECTS FOR FUTURE RESEARCH As in other fields, behavioral neuroendocrinology is experiencing exciting technological developments that promise to significantly increase our knowledge in this area within the next few years. These include developments in molecular biology, genetics, and brain imaging.

5.1. In-vivo Imaging of Brain Activity As we have seen, identifying the brain regions implicated in the regulation of specific behavioral patterns is a fundamental step for understanding their endocrine regulation. In the last 20 years, noninvasive imaging techniques such as positron emission tomography (PET) and functional magnetic resonance imaging (fMRI) have been applied in humans and primates in order to identify brain areas underlying the expression of particular behaviors, and have revolutionized our understanding of brain function (for a review see Otte & Halsband, 2006). The development of scans that fit small animals has

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allowed extension of this technology to other mammalian and nonmammalian species (for a review see Van der Linden, Van Camp, Ramos-Cabrer, & Hoehn, 2007). Functional magnetic resonance imaging has already been applied to fishes (Van den Burgh et al., 1999) but behaviorally relevant questions have not been addressed. It should be pointed out that fMRI requires the immobilization of the animal, limiting the questions that can be tested. Nevertheless, as in humans and primates, brain activation by external cues (e.g., pheromones, sight of a mating partner) in immobilized animals may provide interesting data on the brain areas activated by different social contexts. Endocrine manipulations of immobilized animals exposed to social stimuli could also provide insights into the roles played by hormones in the modulation of brain activity. Manganese-enhanced MRI techniques also can be applied to the study of brain activation in nonimmobilized animals. Manganese enters the cells during voltage-gated Ca2þ channel opening and has a low clearance time, and can thus be used as a contrast agent in MRI to assess brain electrical activity. By administering Mn2þ to animals before the behaviors are elicited and performing MRI scans immediately afterwards it is possible to visualize the brain areas that have just been active. Nevertheless, as in immunohistochemical or in-situ hybridization studies using immediate early gene expression for the same purposes, manganese-enhanced MRI also lacks specificity (see Van der Linden et al., 2007 for details). Combining fMRI and manganese-enhanced MRI with more classical techniques (electrophysiology, lesions, immunohistochemistry, immunoneutralization) seems a promising strategy for identifying the neural substrates underlying the expression of specific behaviors in fishes.

5.2. Molecular Biology Molecular biological tools are valuable technical resources when trying to understand the mechanisms through which hormones change cells and ultimately behavior. There are aspects of hormonal action in relation to behavior for which molecular biological tools are particularly valuable. These techniques can be applied to understanding how changes in gene expression and ultimately protein production influence behavior, and how hormones might mediate these changes. They may also help us to understand how the same hormone may have very different effects in different individuals of the same species or even in the same animal at different times. Hormonal action is influenced by a multitude of factors, including the action of transport proteins, sensitivity of the target tissues to hormones, and interaction with other modulators (neuropeptides, enzymes, other hormones), and molecular biological tools may be useful in characterizing some of these aspects, providing a more integrative view of hormonal action.

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Currently, knowledge of the species genome/transcriptome is growing at an exponential rate, due mostly to new massive sequencing techniques (Morozova & Marra, 2008). This knowledge facilitates the design of methods for gene-expression profiling (e.g., microarrays, RNAseq) and also the interpretation of results from gene-expression studies. These recent advances have allowed documentation of differences in brain gene expression profiles in animals with divergent sexual behaviors (e.g., males and females, dominants and subordinates, bourgeois and sneaker males) (see St-Cyr & Aubin-Horth, 2009 for a list of studies). It seems likely that most of these differences are under hormonal regulation and that some will be behaviorally relevant. Fish behavioral genomics is therefore a growing field and data for non-model species will certainly accumulate over the next few years, allowing comparative functional genomic studies that may provide important insights into the neuroendocrine mechanisms mediating behavior. Nevertheless, knowledge of protein function is clearly falling behind DNA/RNA sequencing, and further effort will be necessary in this respect.

5.3. Genetics New genetic approaches applied to fish behavior are also very promising. There is now a multitude of techniques available for manipulating gene expression in many animals, including fishes, and some recent advances are worth noting. A landmark of gene manipulation techniques in fishes was the development of the first transgenic teleost in 1984 by Zhu, Li, He, and Chen (1985), where a recombinant human growth hormone gene was introduced into a goldfish genome. Since then, many stable transgenic lines have been produced in species such as zebrafish, medaka, carp, and salmon. In 1999, the first zebrafish GAL4-UAS stable transgenic line was produced (Sheer & Campos-Ortega, 1999). In the GAL4-UAS transgenic binary system, the gene encoding a yeast transcription activator protein (Gal4) is placed under the control of a native gene promoter, or driver gene. In another line, a short sequence of the promoter region activated by Gal4 (UAS) is placed upstream of a target gene. By crossing both lines, it is possible to obtain transgenics where the activation of the target gene by Gal4 binding to UAS is restricted to cells expressing the driver gene (i.e., synthesizing Gal4). This system is now highly refined in Drosophila and has been used with enormous success in the study of the neural control of this genus’ behavior, including the neuroendocrine regulation of sexual behavior (e.g., Certel, Savella, Schlegel, & Kravitz, 2007). In zebrafish, the system is not as refined, but recent advances have been made. As an example, in a 2008 study this


system was used in zebrafish to assess the role played by distinct populations of neurons in a specific behavior, the toucheresponse behavior. Gene and enhancer trap methods were first applied to generate fish expressing Gal4 in specific cells. Gal4 gene expression was visualized by crossing Gal4 transgenics with another transgenic line carrying the green or red fluorescent protein (GFP, RFP) gene downstream of the Gal4 recognition sequence (UAS). Finally, parental lines where Gal4 was expressed in relevant cell types were crossed with another transgenic line carrying a synaptic transmission blocker gene (the tetanus toxin light chain gene) downstream of UAS. Gal4 expressed in different neural populations, and thus blocking synaptic transmission in different brain regions, caused distinct abnormalities in the toucheresponse behavior (for details see Asakawa et al., 2008). Transgenic Gal4 medaka lines combined with a heat-shock promoter that allows a temporal control by temperature of the expression of the gene downstream to UAS have also been developed (Grabher & Wittbrodt, 2004). These examples illustrate the potential of genetic manipulation for studying behavior in fishes. The field of behavioral neuroendocrinology will certainly benefit from this technology in the near future.

ACKNOWLEDGEMENTS The writing of this review was partially supported by the following FCT research grants: UI&D 331/2001; PTDC/MAR/71351/2006; PTDC/MAR/69749/2006; PTDC/MAR/72117/2006.

ABBREVIATIONS 11-KA 11-KT 17,20b,21P 17,20bP 17,20aP ACTH ART AVP AVT CNS CYP19 E2 EOD fMRI FSH GFP GnRH GTH HPG HPLC IP IST

11-ketoandrostenedione 11-ketotestosterone 17a,20b,21-trihydroxy-4-pregnen-3-one 17,20b-dihydroxy-4-pregen-3-one 17,20a-dihydroxy-4-pregen-3-one Corticotropin Alternative reproductive tactic Arginine vasopressin Arginine vasotocin Central nervous system See P450aro Estradiol Electric organ discharge Functional magnetic resonance imaging Follicle-stimulating hormone Green fluorescent protein Gonadotropin-releasing hormone Gonadotropin Hypothalamicepituitaryegonadal High pressure liquid chromatography Initial phase Isotocin

Chapter | 7

LH mPOA MT OHT OXY P450aro PET PG PGF2a Pn POA PPn RFP SPPn T TP VAC VPG


Luteinizing hormone Medial preoptic area Methyltestosterone 11b-hydroxytestosterone Oxytocin P450 enzyme aromatase Positron emission tomography Prostaglandin Prostaglandin F2a Pacemaker nucleus Preoptic area Prepacemaker nucleus Red fluorescent protein Sublemniscal prepacemaker nucleus Testosterone Terminal phase Vocaleacoustic integration center Vocal hindbrain-spinal pattern generator

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Chapter 8

Neuroendocrine Regulation in Sex-changing Fishes Earl T. Larson Northeastern University, Boston, MA, USA

SUMMARY Sex change in fishes is a topic that has invoked much interest among physiologists and ethologists alike. At least 14 fish families contain protogynous species, with 11 of these families being common to coral reefs. Sex change can be from female to male (protogyny) or male to female (protandry). Social cues mediate the change process, but these cues trigger a cascade of events in the hypothalamicepituitaryegonadal (HPG) axis. The end result is a total reconfiguration of the gonad and behavior. Chemical signals ranging from classical neurotransmitters to neuropeptides to steroids are involved. This chapter discusses various types of sex reversal, social control, and evolutionary aspects, and most importantly the neuroendocrine control mechanisms of sex reversal in fishes.

1. INTRODUCTION Sex change in fishes is surely one of the most fascinating aspects in the field of vertebrate reproduction. To the layman, the idea is most likely mind-boggling. To the initiated researcher, the idea presents a plethora of interesting research opportunities. Teleost fishes are already fascinating, with the greatest diversity of reproductive strategies of any vertebrate class. Teleosts exhibit internal and external fertilization, oviparity, and viviparity. The variety of reproductive modes includes such diverse strategies as sex reversal, simultaneous hermaphroditism, parthenogenesis, sexual parasitism, and self-fertilization (Breder & Rosen, 1966). The strategy that is most common throughout the animal kingdomdgonochory (separate sexes)dis also abundant in teleosts. While all these modes of reproduction are worthy of discussion, this chapter will tackle the neuroendocrine regulation of sex reversal in teleost fishes. Sex reversal is a complex integrative, multilevel phenomenon that covers multiple aspects of multiple endocrine axes. There are few parts of the endocrine system Hormones and Reproduction of Vertebrates, Volume 1dFishes Copyright Ó 2011 Elsevier Inc. All rights reserved.

that are not involved in sex reversal and we have just begun to uncover the tip of the iceberg when it comes to understanding this process. We are also only beginning to understand the complex social and neurological control mechanisms of sex reversal. This chapter will attempt to summarize the large existing body of work and suggest areas that should be topics for future investigation.

2. HERMAPHRODITISM IN FISHES Sex reversal is a type of hermaphroditism. Hermaphroditism simply refers to a situation in which an organism is capable of reproducing as both a male and a female in the same life history. The simplest form of hermaphroditism is simultaneous hermaphroditism, wherein the individual simultaneously possesses both functional ovarian and testicular tissue (Atz, 1964; Warner, 1984). Simultaneous hermaphroditism is common in the seabasses of the subfamily Serranidae (Sadovy & Domeier, 2005). Different from simultaneous hermaphroditism is sequential hermaphroditism, wherein an individual first possesses functional ovarian tissue and then functional testicular tissue or vice versa. It is sequential hermaphroditism that will be the focus of this chapter. Sequential hermaphroditism appears in two different forms. Protogyny (‘female first’) is the most common type of sequential hermaphroditism in teleosts. In this strategy, an individual matures sexually as a female and then becomes male later in the life history. Protandry (‘male first’) is a strategy in which individuals mature as males and later become female. Although sex reversal can occur before an animal reaches sexual maturity (Shapiro, 1988), it occurs more commonly after maturity. On coral reefs, there are many hermaphroditic fishes (Smith, 1982). At least 14 fish families contain protogynous species (see Table 8.1); 11 of these families are common on coral reefs (Warner, 1984). In the families Labridae (wrasses), Scaridae (parrotfishes), and Serranidae 149

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TABLE 8.1 Modes of sex change in 15 families. Representative genera are listed for each family and each mode Family

Monadric protogynous

Diandric protogynous

Tenualosa 2

Emmelichthyidae (rovers)

Spicara

Gobiidae (Gobies)

Coryphopterus3, Gobisoma4

Trimma5, Lythrypnus6 Gonostoma7, Cyclothone8

Gonostomatidae (bristlemouths) Berggylta9, Bodianus10, Clepticus10, Labroides11, Semicossyphus12

Halichoeres10, Thalassoma13, 14 Lethrinus15

Lethrinidae (emperors) Muraenidae (moray eels)

Bidirectional

1

Clupeidae (herrings)

Labridae (wrasses)

Protoandrous

Muraena, Gynothorax, Echinda, Uropterygius16 Inegocia17

Platychephalidae (flatheads) Pomacanthidae (angelfishes)

Centropyge18, Genicanthus19

Pomacentridae (damselfishes)

Dascyllus20

Amphiprion21 Pseudochromis22

Pseudochromidae (dottybacks) Scaridae (parrotfishes)

Calotomus, Cryptotomus, Leptoscarus, Sparisoma23

Serranidae (seabasses)

Epinephalus24, Mycteroperca25, Pseudanthias26

Sparidae (porgies)

Chrysoblephus27, Pagrus28

Synbranchidae (rice eels)

Monopterus31

Calotomus, Cetoscarus, Scarus, Sparisoma23

Acanthopagrus29, Rhabdosargus30 Synbranchus32

1

17

2

18

Blaber et al. (1996). Dulcic, Kraljevic, Grbec, and Cetinic (2000). 3 Cole and Shapiro (1990). 4 Robertson and Justines (1982). 5 Sunobe and Nakazono (1993). 6 St. Mary (1993). 7 Badcock (1986). 8 Miya and Nemoto (1985). 9 Elofsson, Winberg, and Nilsson (1999). 10 Warner and Robertson (1978). 11 Robertson (1972). 12 Cowen (1990). 13 Warner and Hoffman (1980). 14 Ross (1983). 15 Grandcourt (2002). 16 Fishelson (1992).

(seabasses), protogyny is nearly exclusive. Protandry occurs in eight fish families, three of which are common on coral reefs. Perhaps the best-known example of protandrous fishes are the anemone fishes of the genus Amphiprion (family Pomacentridae) (Warner, 1984). Simultaneous hermaphroditism is most common in deep-sea fishes that infrequently encounter conspecifics, and one sub-family of the seabassesdSerraniae, composed primarily of simultaneous hermaphroditesdis restricted to coral reefs.

Shinomiya, Yamada, and Sunobe (2003). Lutnesky (1996). 19 Shen and Liu (1976). 20 Moyer and Nakazono (1978). 21 Godwin (1995). 22 Wittenrich and Munday (2005). 23 Streelman, Alfaro, Weastneat, Bellwood, and Karl (2002). 24 Shapiro and Sadovy (1993). 25 Coleman, Koenig, and Collins (1996). 26 Shapiro (1981). 27 Garrat (1986). 28 Pajuelo and Lorenzo (1996). 29 Hesp, Potter, and Hall (2004). 30 Yeung and Chen (1987). 31 Liem (1963). 32 Lo Nostro and Guerrero (1996).

Early research on hermaphroditic fishes consisted of compilations of hermaphroditic species. These descriptions were based mainly on gonadal histology (Shapiro, 1988). During the 1970s, more behavioral and manipulative work was carried out both in aquaria and in the field on a variety of coral reef species to characterize behaviors associated with sex reversal. Most of this work concentrated on three genera: the diandric protogynous labrid (Thalassoma bifasciatum) (Reinboth, 1975; Warner & Hoffman, 1980),

Chapter | 8

Neuroendocrine Regulation in Sex-changing Fishes

the monandric protogynous serranid (Pseudanthias squamipinnis) (Fishelson, 1970; Shapiro, 1981), and the protandrous pomacentrids (Amphiprion spp.) (Moyer & Nakazono, 1978; Fricke, 1979). During the 1980s, work on Thalassoma duperrey suggested that different mating systems may exist in closely related species and that this may produce differences in proximate control of sex reversal (Ross, 1986). Basic behavioral work has continued and recently has brought to light new and exciting discoveries.

2.1. Protogynous Sex Reversal In protogynous species, females usually outnumber males in a population (Warner, 1984; Sadovy & Shapiro, 1987), and differences exist in the mating behaviors of the two sexes. Additionally, there are coloration differences between the two sexes, commonly referred to as dichromatism. There are two types of protogyny: monandric and diandric. In monandric protogyny, all males in the population are derived from females. In diandric protogyny, individuals begin life as a male or a female in the initial color phase, which is female-typical coloration. Later the initial phase (IP) male or female may change into a terminal phase (TP) male. Terminal phase males exhibit a maletypical coloration pattern not seen in females or IP males, known as TP coloration. Initial phase males also are called primary phase males or nonterminal males. The three possible phenotypes in a diandric system exhibit three different reproductive behaviors. Terminal phase males dominate spawning and hold territories, which they defend using aggressive displays. They will pair spawn with multiple females each day. Females usually travel within the territories, and will spawn once per day. Initial phase males do not hold territories, are subordinate, and only mate by sneaking copulation (Warner, 1984). The number of IP males present in a population increases with the size of the population (Warner, 1984; Ross, Hourigan, Lutnesky, & Singh, 1990). The only chance for IP males to mate is by streaking or group spawning. In streaking (also called sneak spawning), a nonterminal male swims into the territory of a TP male when the TP male is pair spawning with a female. When the female releases eggs, the IP male quickly swims by and releases milt. Competition takes place between the sperm of the TP and IP males. Group spawning takes place when more than two IP males swarm a female until she releases her eggs. All the IP males present then attempt to fertilize the eggs. Both streaking and group spawning result in sperm competition. Initial phase males usually have testes that are larger and capable of greater sperm production than TP males (Hourigan, Nakamura, Nagahama, Yamauchi, & Grau, 1991). The saddleback wrasse (T. duperrey), the most common species of wrasse inhabiting coral reefs of the Hawaiian Islands (Hourigan & Reese, 1987), is one of many

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protogynous coral reef species (Ross, 1986). T. duperrey is diandric, but the social organization is slightly different from most types of diandric wrasses in that females have a larger home range and do not always mate with the same male. Ross (1986) described the mating system of T. duperrey as a resource defense polygyny: a mating system wherein a male defends a territory and mates with multiple females within that territory. Terminal phase males are large with relatively small testes, and they spawn individually with females in their spawning territories. Initial phase males are smaller with relatively large testes and group spawn in a large group of females and IP males (Hourigan et al., 1991). Sex reversal is controlled by the ratio of larger (usually male) to smaller (usually female) fish in the population (Ross, Losey, & Diamond, 1983). Using a series of visual, tactile, and chemical barriers, it has been demonstrated that the cues responsible for the initiation of sex reversal are visual and not chemical or tactile. The presence of smaller individuals promotes sex reversal and the presence of larger individuals inhibits sex reversal. Therefore, if the ratio of other individuals in the population is heavily weighted toward smaller fish, a female will undergo sex reversal, thus increasing her fitness (Warner, 1984). Morrey, Nagahama, and Grau (2002), however showed that, in T. duperrey, it was not size, but color phase, that mattered. If a female of the appropriate size to change sex was in the presence of a TP male, she would not change sex, even if that male was larger than her. Conversely, she would change sex in the presence of an individual with IP coloration regardless of whether the individual was female or male (Morrey et al. 2002).

2.2. Protandrous Sex Reversal The best-characterized examples of protandry have been in the anemonefishes, which are the subfamily Amphiprioninae belonging to the family Pomacentridae, or damselfishes (Moyer & Nakazono, 1978). In the genus Amphiprion, the populations usually consist of small groups (fewer than 10) of fish that are associated with a home anemone. There is a monogamous pair of sexually mature adults, one secondary female and one primary male (Moyer & Nakazono, 1978). There is no sexual dichromatism in anemone fishes, but the female is always the largest individual in the population. The second largest fish is a functional male. The other members of the population are sexually undifferentiated. If the female disappears, the male will undergo sex reversal and become female. At this point, the largest of the undifferentiated individuals will mature as a male, which will also happen if the male disappears but the female does not. In the Sparidae (porgies), protandry has been characterized by histology (Sadovy & Shapiro, 1987; Hesp, Potter, & Hall, 2004). Protandrous porgies begin life as males, but they have an ovotestis that is predominated by

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testicular tissue. During the first season of reproduction, this testicular tissue actively produces sperm. At the end of the season the testicular tissue regresses. In subsequent seasons, the testicular tissue can again develop and produce sperm or the ovarian tissue can develop and produce eggs (Hesp et al. 2004). Once an individual has become a functional female, it will continue to be female for the rest of its life (Hesp et al. 2004). Little is known about the control of protandrous sex change in the sparids.

2.3. Bidirectional (Serial) Sex Reversal In protogynous and protandrous species, sex change only happens once. The goby Trimma okinawae (family Gobiidae) was described as the first known teleost capable of serial sex change (Sunobe & Nakazono, 1993). Individuals begin life as females and can change to males upon removal of the dominant male. Unlike labrids, however, this goby retains ovarian tissue within the functional testes. If a larger, more dominant male appears, the newly transformed male will change back to a female (Sunobe & Nakazono, 1993). Since Sunobe and Nakazono’s work, other gobies have been shown to be capable of bidirectional sex reversal. Grober and colleagues have conducted the most extensive examinations of bidirectional sex change, using the goby Lythripnis dalli (Perry & Grober, 2003). Bidirectionality in this species was first described by St. Mary (1993), and the bidirectional switch is under social control (Rodgers, Drane, and Grober, 2005). Initially, they showed that protogynous sex reversal is mediated by social cues and can be induced by removing males (Reavis & Grober, 1999). This goby lives in small social groups of 4e12 individuals wherein one is usually male and each group is usually associated with a sea urchin (Centrostephanus coronatus) (Lorenzi, Early, & Grober, 2006). When investigators constructed all-female groups, the dominant female would change sex to male, and when they constructed all-male groups, the subordinate males would change sex to females (Rodgers, Earley, & Grober, 2007). This led to the description of a simple operating principle: if subordinate, one expresses femaleness; if dominant or not subordinate, one expresses maleness (Rodgers et al., 2007). These results suggest that an individual will recognize its place in a dominance hierarchy, and then exhibit a sexual phenotype accordingly. This is not the first time that fishes have been shown to recognize their position in a dominance hierarchy (Winberg, Nilsson, & Olsen, 1991; Fox, White, Kao, and Fernald, 1997; Leiser & Itzkowitz, 2004; Sneddon, Margareto, & Cossins, 2005), but it is among the better demonstrations of such in sex-changing fishes. Additional experiments have determined that behavioral interactions are the most efficient factors controlling sex change in L. dalli, though visual and olfactory cues have an influence


as well (Lorenzi et al., 2006). Although the majority of the work has been done in the laboratory, important field studies have indicated that sex reversal demonstrated in the laboratory also occurs in the field but at a slower rate (Black et al., 2005). Additional field work has demonstrated that there are two different male phenotypes in L. dalli (Drilling & Grober, 2005). There is an alternative male phenotype, dubbed mini-males by the researchers. These mini-males are similar to most IP males of other diandric species in that they have larger gonads adapted for greater sperm production. The difference is, however, that in L. dalli there are no initial and terminal color phases; the coloring is the same regardless of sexual phenotype (Drilling & Grober, 2005).

3. HYPOTHESES OF NATURAL SEX REVERSAL While most of this chapter will focus on the proximate mechanisms of sex reversal, ultimate mechanisms will first be considered. Sequential hermaphroditism, or in fact any type of hermaphroditism, is largely a question of sex allocation. Sex allocation theory is an important question in evolution (Charnov, 1982). Probably the most important part of sex allocation theory as it relates to sequential hermaphroditism is Ghiselin’s (1969) size advantage hypothesis (SAH). The hypothesis states that, if an individual reproduces more efficiently as one sex when smaller and another sex when larger, then sex change is an advantageous strategy. In protogynous species, large males typically monopolize mating events (Warner, 1984). In these situations, smaller males have difficulty spawning, but smaller females do not. In this size-dependent situation, it would be advantageous for an individual to be female when small and male when large. Protandry is advantageous when monopolization does not occur and mating consists of random pairing. This is the case for many sparids (Buxton & Garratt, 1990). Perhaps the best known case of protandry is in anemonefishes (Amphiprion spp.) (Shapiro, 1992). In this genus, populations are usually very small and mating is random. Therefore it is most advantageous for the largest animal to be a female so that more eggs, the limiting gamete, are produced. If fertility is plotted against size for each sex, it is the point where these lines intersect that should indicate when it is most advantageous for an animal to change sex (Figure 8.1). Warner (1988), however, after factoring in growth and mortality, suggested that it is the reproductive value (RV) (the expected future reproductive success) that is more important than just fertility. Sex change should be favored when (1) the RV is closely related to age or size and (2) the relationship differs between the sexes. In this case, selection favors a strategy to first be the sex where RV increases more slowly and then change to the other sex at a later stage (Allsop & West, 2003).

Chapter | 8

(a)


(b)

FIGURE 8.1 The size advantage hypothesis. Female reproductive value is represented by the slightly upward curving line in both graphs and depicts that reproductive value (RV) is directly related to body size because the number of eggs produced is directly related to body size. (a) Protogyny. In situations that favor protogyny, small males do not hold territories and have to rely on sneak or group spawning, so RV is low. As they grow, males get to the point where they can defend a territory; it is at this point that RV increases dramatically. The point at which the male and female lines intersect is the best time for the female to become male. (b) Protandry. The anemone fish is a good example of this situation. There is a small group of fish living on the anemone with one reproductive female, one reproductive male, and several nonreproductive juveniles. The male’s RV remains constant because he only has one female with which to reproduce. The female’s RV increases with body size. The point at which the lines intersect is the best time for the male to change to female. Reproduced from Munday, Buston, and Warner (2006).

Reproductive value seems to be more useful in explaining why some individuals change at different sizes (Munday, Buston, & Warner, 2006). The phenomenon of early sex change has been discussed for quite some time. Early sex change is typically when a female changes sex at a size too small to be a breeding male. This could be a viable strategy if nonreproductives have decreased mortality or increased growth (Munday et al., 2006). What actually makes the sex change ‘early,’ however? It would not be unreasonable to think that, as with most other phenotypic traits, variation might exist in the size of an individual at the time of sex change. Variation in body size at the time of sex change has been discussed almost as much as ‘early sex change.’ One of the most interesting studies was conducted by Allsop and West (2003), who looked at 61 populations of 52 sex-changing species, including protandrous as well as both monandric and diandric protogynous species. Allsop and West (2003) added empirical measurements to test earlier theoretical work (Charnov 1982; Charnov, Berrigan, & Shine, 1993; Charnov & Sku´lado´ttir, 2000). They built upon a model called ‘dimensionless life history theory’ (Charnov & Sku´lado´ttir, 2000). Using several dimensionless properties such as mortality rate, age at first breeding, and the relationship between male fertility and size, Charnov and Sku´lado´ttir (2000) predicted that populations or species that have similar values for these dimensionless traits will have similar values for (1) size at sex change/maximal size, (2) age at sex change/age at first breeding, and (3) proportion of the breeding population that is male (Allsop & West, 2003). After assembling the data from the 52

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different species, Allsop and West (2003) tested the model and found that, for the 52 species tested, fishes change sex at a constant proportion of their maximal sizedthat proportion being 0.79. This means that, in general, fishes change sex when they reach a length that is 79% of their maximal length. The authors also found that fishes change sex when the age of the sex change divided by age of maturity equals 2.5 (Allsop & West, 2003). That is, most fishes change sex when they are 2.5 times as old as when they sexually matured. Therefore, the combination of data on size and RV is thought to provide a stronger indication of when it is most advantageous for an individual to change sex than does the size advantage hypothesis.

4. SOCIAL FACTORS AFFECTING SEX REVERSAL In almost all documented cases, sequential hermaphroditism is under social control (Warner, 1984; Perry & Grober, 2003; Munday et al., 2006). The phenomenon of social control of sex reversal was first reported independently in the early 1970s for two different species: the fairy basslet (P. (Anthias) squampinnis) (Fishelson, 1970) and the cleaner wrasse (Labroides dimidiatus) (Robertson, 1972). Since then, social control has been documented for most of the families mentioned in Table 8.1, including Gobiidae, Labridae, Pomacentridae, Scaridae, and Serranidae. This social control is often referred to as behavioral sex determination (BSD), which is considered to be a form of environmental sex determination (ESD) (see Chapter 1, this volume). In this case, it is the social environment rather than the physical environment (e.g., temperature) that determines sex. If one looks throughout the animal kingdom, these two modes are seen much less frequently than genetic sex determination, which is the mode used in most vertebrates. On coral reefs, however, there is no shortage of BSD among teleosts. Two initial studies demonstrated that, by removing the male in a group, a female will change sex (Fishelson, 1970; Robertson, 1972). These studies opened up a new field of research. Initially researchers studied the ethology of sex change and the social control and then expanded to physiological mechanisms, which will be discussed later. When a female (or IP male) experiences a social cue that initiates transformation to a TP male, the first changes are behavioral, and these can take place the same day (Warner & Swearer, 1991). Various studies have given detailed accounts of changes in behavior concomitant with protogynous sex reversal. Shapiro (1981) reported an increase in male-associated behaviors, especially territorial behaviors in P. squamipinnis. Nose rushes, dorsal-spine erections, and other aggressive behaviors increased in both number and frequency. In wrasses, Warner and Swearer (1991) found that, within minutes of the removal of the

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TP male from a patch reef, the largest female present exhibits aggression toward other large females and courtship towards smaller females. The aggressive behaviors are thought to inhibit other females from changing sex (Robertson, 1972). Dominance hierarchies likely play a role here. While it seems to be the largest female that is changing sex, it could well be that, as well as being largest, she is most dominant. Dominance hierarchies, commonly known as pecking orders, are common in fishes (Winberg et al., 1991; Shartl et al., 1993; Reichard et al., 2005; Henningsen, Murru, Rasmussen, Whitaker, & Violetta, 2008). These social structures could be important in determining which individual changes sex.

5. NEUROENDOCRINE FACTORS AFFECTING SEX REVERSAL The role of the hypothalamicepituitaryegonadal (HPG) axis in sex reversal is undeniable. Despite this, there are many questions about the role of the different components of the axis that remain unanswered. Are steroids important in the control of sex reversal or are they merely the end product of the process? Is the hypothalamus the control switch for the process? If so, what are the control signals: neuropeptides, classical neurotransmitters, or both? Initially, all work on how the HPG axis might be involved in sex change was based on histological observations (see review by Shapiro, 1988). Following these initial histological characterizations, attention shifted to the physiological mechanisms underlying sex reversal. Research usually involved injecting different substances into IP individuals and observing resultant color or histological changes as females transformed into males (Stoll, 1955; Reinboth, 1975). The intention of the researchers was to select females for the injections, but selection was based on coloration (hence IP individuals) and not gonadal morphology, which was impossible to determine without sacrificing the animals. The majority were likely females, based on the percentage of IP animals that are female, but the occasional IP male cannot be ruled out.

5.1. Gonadal Steroids The first of these studies focused on androgens (Stoll, 1955; Reinboth, 1975). These early studies provided inconsistent results (Shapiro, 1988). Whereas androgens appeared to initiate sex reversal in some species, results were less convincing in others (see Shapiro, 1988). Exogenous androgens had been used extensively in aquaculture to induce sex reversal and increase growth in gonochoristic species such as Ictalurus punctatus and Oeochromis mossambicus (Gannam & Lovell, 1991; Kuwaye et al., 1993). Various laboratories injected androgens into females of different protogynous species to induce sex reversal


(Kramer, Koulish, & Bertracchi, 1988; Cardwell & Liley, 1991). In the only study to follow protogynous animals after the withdrawal of exogenous androgens, Tan-Fermin (1992) found that the grouper (Epinephelus suillus) reverts back to female four months after withdrawal of 17-a methyltestosterone treatment. A parallel line of research that began after successful induction of sex reversal via exogenous factors was the characterization of endocrine changes throughout the natural process of sex reversal. Although it is still unclear what role steroids serve in the process of sex reversal, they do play a role. It is uncertain, however, whether that role is driving the change or is a result of the change. If changes in sex steroids are a result of the sex change and not driving the reversal, they could feed back on the upstream regulators, such as peptides or monoamines, which might be driving the process to signal completion of sex reversal. One thing is clear: steroid levels change over the course of sex reversal. This is seen in protandrous, protogynous, and bidirectional species. In protogynous fishes, which are the most studied, we see a general decrease in estrogens and increase in androgens throughout the process (Nakamura, Hourigan, Yamauchi, Nagahama, & Grau, 1989). In protandrous fishes, the opposite happens: androgens decrease and estrogens increase (Godwin, 1993). In bidirectional fishes androgens do not change with reversal, but estrogens increase as individuals become female and decrease as they become male (Lorenzi, Early, Rodgers, Pepper, & Grober, 2008). With the development of radioimmunoassay by Yalow and Berson (1959), measurement of gonadal steroids in fishes, including sex-changing fishes, became a possibility. Some of the first studies to investigate changes in steroids accompanying sex reversal in natural populations were done on the saddleback wrasse (T. duperrey) and the stoplight parrotfish (Sparisoma viride) (Nakamura et al., 1989; Cardwell & Liley, 1991). Female saddleback wrasse have higher circulating levels of estradiol (E2) than do TP males that are derived from females (Nakamura et al., 1989). Terminal phase males have higher levels of 11-ketotestosterone (11-KT) than do females. It is important to take note of the fact that teleost fishes have high levels of both 11-KT and testosterone (T) in the bloodstream. It is 11-KT that seems to be the more active androgen in male teleosts and is the hormone that shows more marked differences in concentrations between males and females. The diandric saddleback wrasse has two male phenotypes: TP males that are derived from females via sex reversal, and IP males that externally have the same coloration as females but are born male and reproduce as males at a smaller size via group spawning or streak spawning (Ross et al., 1990). Hourigan et al. (1991) discovered that sperm production is much higher in IP than

Chapter | 8


TP males of this species and that, when expressed as a function of body size, the testes are five to six times larger that those of TP males. Using an in-vitro approach, Hourigan et al. (1991) looked at the ability of testicular tissues from IP and TP males to produce T and 11-KT when provided with 14C-labeled steroidal precursors such as 17aOH-progesterone, dehydroepiandrosterone (DHEA), or T and exposed to gonadotropin (GTH). Terminal phase males, while having much smaller gonads, have steroidproducing Leydig cells that are both greater in number and better developed as compared to those of IP males (Figure 8.2) (Hourigan et al. 1991). Testes of TP fish are also more efficient at producing 11-KT and T (Hourigan et al. 1991). On the other hand, IP males are very proficient at producing sperm. Their main mode of reproduction is

FIGURE 8.2 Leydig cells in the testes of (a) initial phase (IP) and (b) terminal phase (TP) males of Thalassoma duperrey. Terminal phase males have steroid-producing Leydig cells (L) that are both greater in number and better developed as compared to those of IP males. Magnifications are 8700 and 7830, respectively. e, erythrocytes in capillaries; s1, primary spermatocyte. Reproduced from Hourigan, Nakamura, Nagahama, Yamauchi, and Grau (1991).

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group spawning (Ross, 1986), which sets up a case for sperm competition. Therefore, the males must produce and release more sperm to have a better chance at successful fertilization. It has been shown in T. bifasciatum that groupspawning males release six times more sperm on average than pair-spawning males (Shapiro, Marconato, & Yoshikawa, 1994). This correlates nicely with the measurement in T. duperrey of IP males having testes five to six times larger than their TP counterparts. How then are the IP males producing so much sperm when they produce so little 11-KT? Cardwell and Liley (1991) conducted a comprehensive study on the diandric stoplight parrotfish (S. viride) wherein they compared steroid levels in (TP) males, transitional individuals, and females. They examined three steroids: 11-KT, T, and E2. As sex change in this species is coupled with color (phase) change, the authors considered that the steroids also play a role in the color change. Females had high levels of E2, lower levels of T, and very low levels of 11-KT (Figure 8.3). Testosterone is present in females as the precursor to E2. As female fish underwent sex and color change, there was a decrease in circulating E2 and an increase in 11-KT, while T remained largely unaffected (Figure 8.3). Treating functional adult females with 11-KT caused sex change and color change (Cardwell & Liley, 1991). As this species is diandric, the authors also measured steroid levels in the IP males. Somewhat surprisingly, these individuals had very low levels of 11-KT and moderate levels of T and E2, even though IP males have large functional testes and are able to spawn (Cardwell & Liley, 1991). In both S. viride and T. duperrey, it seems possible to have spermatogenesis without high levels of 11-KT. Treatment with T can drive spermatogenesis in other species although T is less potent than 11-KT (Norris, 1987). In the lungfish, 11-KT is nondetectable and T is correlated with spermatogenesis (Joss, Edwards, & Kime, 1996). Cardwell and Lily (1991), however, conclude that 11-KT plays an important role in the initiation of sex and color change in S. viride. In the first work to chronicle changes in steroids with sex change in a protandrous species, Godwin and Thomas (1993) examined an anemonefish (Amphiprion melanopus). They found that males have higher levels of 11-KT but lower levels of T, E2, and androstenedione (AND) (a precursor of both T and E2) as compared with females. Unlike other fishes, there is not a gradual change in the plasma levels of these steroids throughout the process (Godwin & Thomas, 1993). Instead, different patterns are seen for each steroid. For example, 11-KT stays high throughout the process, dropping only when the animal has completed the transition to female. Likewise, estrogens stay low until the animal completes the change to female, and the same is true for AND. The most interesting pattern occurs in T, which starts at intermediate levels in males,

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(a)

(b)

(c)

Marsh, Creutz, Hawkins, & Godwin, 2006). In the gonads of the serial sex-changer T. okinawae, aromatase seems to be important for the development of oocytes when an individual is changing from male to female (Sunobe et al., 2005). Additionally, there are two isoforms of aromatase, cyp19a1 and cyp19a2. The former is expressed in the gonad and the latter in the brain (Gardner et al., 2005). These two isoforms of aromatase are by no means restricted to sex-changing fishes, and have been reported in other species including the goldfish and the zebrafish (Tchoudakova & Callard, 1998; Trant, Gavasso, Ackers, Chung, & Place, 2001). In the protandrous black porgy (Acanthropargus schlegeli) dual roles have been observed for cyp19a1 (Wu, Tomy, Nakamura, & Chang, 2008). This fish undergoes male sexual differentiation during the juvenile stage and possesses bisexual gonads during the first two years of its life. These bisexual gonads change to functional ovaries at three years of age (Wu et al., 2008). During the initial male sexual differentiation, high levels of cyp19a1 occur in the undifferentiated gonad. After testicular maturation, levels of cyp19a1 are low and then increase with sex reversal, reaching high levels again in the resultant females. Therefore, the authors concluded that cyp19a1 has an important dual role in both testicular and ovarian differentiation (Wu et al., 2008).

5.2. Peptides FIGURE 8.3 Plasma steroid levels in male, female, and transitional Sparisoma viride. The values are given as means standard error of the mean (SEM) with the sample sizes in the bars. The top, middle, and lower panels represent 11-ketotestosterone, testosterone, and estradiol, respectively. Reproduced from Cardwell and Liley (1991).

drops to very low levels midtransition, and then increases to high levels in females (Godwin & Thomas, 1993). The authors conclude that 11-KT is important for the maintenance of male function, and estrogens are responsible for female functions. They conclude that there is no special role for AND in sex change and that E2 is resultant of sex change and not causative because an increase in E2 is not seen until the transition to female is complete. One interesting piece of work that ties together sex steroids and GnRH showed that injections of androgens increased the number of GnRH neurons in the hypothalamus (Grober, Jackson, & Bass, 1991). This suggests that there must be some sort of feedback from gonadal steroids that affect the hypothalamus. More recently, neurosteroids have become of interest in the field of sex reversal. Aromatase has been a focus of these studies, which have looked at aromatase expression in both the brain and the gonads (Gardner, Anderson, Place, Dixon, & Elizur, 2005; Sunobe, Nakamura, Y. Kobayashi, T. Kobayashi, & Nagahama, 2005;

Eventually, researchers started to investigate the sexreversal effects of factors that control the release of androgens such as GTHs and gonadotropin releasing hormone (GnRH) (Koulish & Kramer, 1989; Kramer, Caddell, & Bubenheimer-Livolsi, 1993). These results again were mixed and, although GTH alone induced sex reversal, GnRH was capable of this induction only in combination with a dopamine (DA) antagonist. These results suggest that neural factors also may be involved in the initiation and maintenance of sex reversal. Subsequent studies looked at other neural aspects of protogyny. Sex reversal in T. bifasciatum was accompanied by an increase in the number of GnRH immunoreactive (ir) neurons in the preoptic area (POA) of the hypothalamus, an area thought to be associated with male function (Grober et al., 1991). In this species, TP individuals had a greater number of GnRH-ir neurons than did IP animals (Grober & Bass, 1991). It is interesting to note that, in the protandrous anemone fish A. melanopus, males also have greater numbers of GnRH-ir neurons in the POA, but in this genus it is the female that is always the terminal state (Elofsson, Winberg, & Francis, 1997). So it seems that in sex-changing fishes GnRH may be correlated with male function rather than sex reversal itself. This is further supported by the evidence that GnRH alone cannot induce sex reversal (Kramer et al., 1993).

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In addition to the work done on the protandrous anemone fish, Elofsson, Winberg, and Nilsson (1999) completed an investigation of the protogynous ballan wrasse (Labrus bergylta). This species is monandric and is unusual in that it does not live on coral reefs, but rather in the cold waters of the eastern north Atlantic. Being monandric, all males in this species are derived from females. This makes for a system that is less complicated than diandric species, because one knows that all males are derived from males and the researcher can focus on a single developmental pathway. Due to its temperate habitat, the ballan wrasse is a strictly seasonal spawner. During the summer spawning season, each male defends a spawning territory and a harem of between three and seven females (Elofsson et al., 1999). As with other wrasses, removal of the largest male results in the largest female changing sex. Males have a greater number of GnRH neurons in the POA than do females. In males, the size of the GnRH neurons correlates with the spawning cycle, and spawning males have larger neurons. The most interesting aspect of this work, however, is that the authors found that in males, but not in females, the number of GnRH neurons correlates with body size (Elofsson et al., 1999). Since larger males are capable of defending larger territories and maintaining a larger harem, this relationship could relate to dominance and/or aggression. Neuropeptides have a role in sex reversal as well as in the overall complexity of the physiological process of sex reversal. In T. bifasciatum, females have lower levels of arginine vasotocin (AVT) than do males (Godwin, Sawby, Warner, Crews, & Grober, 2000). The goby T. okinawae, one of the few species shown to be capable of multiple sex reversals, shows similar but rapid and reversible changes in the number of AVT-producing neurons with serial sex reversals (Grober & Sunobe, 1996). In the gonochoristic white perch (Morone americana), AVT increases male courting behavior (Salek, Sullivan, & Godwin 2002). Although there are some suggestions that AVT is related to sex change, it may play more of a role in dominance and aggression as reported for zebrafish (Larson, O’Malley, & Melloni, 2005) and rainbow trout (Backstro¨m & Winberg, 2009). In T. bifasciatum (Godwin et al., 2000), males have more AVT mRNA than females. The territorial TP males have three times more AVT mRNA than nonterritorial IP males (Godwin et al., 2000). This correlates with work on a non-sex changing cichlid, Astatotilapia burtoni, a member of the same suborder as wrasses, parrotfishes, and angelfishes, three of the most important sex-changing families. Territorial males had higher levels of AVT mRNA than did nonterritorial males (Greenwood, Wark, Fernald, & Hoffmann, 2008). Additionally, Semsar and Godwin (2004) found that ovariectomized female T. bifasciatum that undergo behavioral sex reversal after male removal (Godwin, Crews, & Warner, 1996) still show increases in

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AVT mRNA. Inseparable from behavioral sex change in this species is the acquisition of territoriality. It is clear that AVT does play a role in territoriality and dominance and possibly in behavioral sex change as well. However, it is clear that AVT does not drive gonadal sex change (Semsar & Godwin, 2004). More work needs to be done to elucidate exactly how much of a contribution AVT makes to behavioral sex reversal vs. territoriality, aggression, and dominance. Since these changes are usually concomitant, this could prove difficult.

5.3. Monoamines Neurobiological questions are an important approach to understanding the phenomenon of sex reversal. When looking for a controlling mechanism for such a complicated process, it is important to address higher-level control. Most studies have focused on downstream events. Although gonadal and pituitary events are, without doubt, crucial to the process of sex reversal, these events are affected by higher levels of control. Hypothalamic peptides are likewise controlled by other neural messengers. An important group of chemicals not previously investigated are the monoamine neurotransmitters or biogenic amines. The role of monoamine neurotransmitters in regulating sex reversal in T. duperrey was investigated by Larson, Norris, Grau, and Summers (2003) and Larson, Norris, and Summers (2003). These amines are important in controlling secretion of hypothalamic and pituitary hormones that previously have been investigated in physiological studies of protogyny and are probably an important part of the pathway mediating conversion of a social cue into a neuroendocrine event. These amines are important in controlling factors that have previously been investigated in physiological studies of protogyny. In gonochoristic fishes, monoamines are known to be the most important neurotransmitters to control secretion of GnRH and GTH, which in turn control secretion of gonadal steroids. Dopamine and a number of compounds known to affect DA release and action have been shown to regulate GTH release in the goldfish (Carassius auratus) (Chang & Peter, 1983a). Dopamine also inhibits GnRH release (Dufour et al., 2005). Intraperitoneal injections of DA and its agonist, apomorphine (APO), decrease serum GTH levels, whereas intraperitoneal administration of the DA antagonists pimozide and metoclopramide has the opposite effect (Peter et al., 1986). Norepinephrine (NE) injected into the third cranial ventricle increases GTH release (Chang & Peter, 1983b) and stimulates GnRH release from brain slices (Yu & Peter, 1992) and GTH release by pituitary fragments in vitro (Chang, Van Goor, & Acharya, 1991). These actions are mediated by a1-like adrenergic receptors. Intraperitoneal injections of serotonin (5-HT) into goldfish also result in

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increased serum GTH levels (Somoza, Yu, & Peter, 1988). Monoamines are linked to reproductive behavior in fishes. Holdway, Sloley, Munkittrick, Downer, and Dixon (1988) have demonstrated that sexually active male flagfish (Jordanella floridae) have elevated 5-HT levels as compared with nonactive males. Dominant males exhibit lower NE levels than subordinate males. In sexually active male Arctic char (Salvelinus alpinus), dominant males have elevated 5-HT levels compared with subordinate males (Elofsson, Mayer, Damsga˚rd, & Winberg, 2000).

6. STUDIES ON THE SADDLEBACK WRASSE The saddleback wrasse (T. duperrey) is one of the beststudied protogynous coral reef species (Ross, 1983; Ross et al., 1983; Ross, 1986, Hoffman & Grau, 1989; Nakamura et al., 1989; Hourigan, et al., 1991; Morrey et al., 2002; Larson et al., 2003a; 2003b). Triggering of gonadal sex reversal is accomplished experimentally by manipulating the ratio of larger (usually male) to smaller (usually female) fish in the population (Ross et al., 1983). However, nothing has been published on the timing of the behavioral changes associated with sex reversal in this species. Ross (1986) described the mating system of T. duperrey as a resource-defense polygyny. As described above, T. duperrey is diandric, but the social organization is slightly different from most types of diandric wrasse in that females have a larger home range and do not always mate with the same male. There are three gender conditions, each with a specific reproductive strategy. Terminal phase males are large with small testes and spawn individually with females in their spawning territories, whereas IP males are smaller with large testes, do not defend mating sites, are subordinate to larger individuals, and mate by either group spawning or streaking. Females can become TP males under the proper social conditions. Sex reversal is controlled by the ratio of larger (usually male) to smaller (usually female) fish in the population (Ross et al., 1983). Using a series of visual, tactile, and chemical barriers, these authors demonstrated that the cues responsible for initiation of sex reversal are visual and not chemical or tactile. The presence of smaller females promotes sex reversal and the presence of TP males inhibits sex reversal. In other wrasse species, when a female experiences a social cue that initiates transformation into a TP male, the first changes are behavioral, and these can appear within hours (Robertson, 1972; Warner & Swearer, 1991). When the TP male bluehead wrasse (T. bifasciatum) was removed from a patch reef, the largest female present began to exhibit aggression toward other large females and to court smaller females within minutes following removal (Warner &


Swearer, 1991). During the mating period the same day, these females exhibited male spawning behavior. By day three following male removal, the largest females had begun the process of color change associated with sex reversal. These behaviors are thought to inhibit other females from changing sex (Robertson, 1972). Shapiro (1981) reported an increase in male-associated behaviors, especially territorial behaviors in P. squamipinnis. Nose rushes, dorsal-spine erections, and other aggressive behaviors increased in both number and frequency. Work done by Ross (1986) on T. duperrey suggests that different mating systems may exist in closely related species and that this may produce differences in proximate control of sex reversal. In T. bifasciatum, sex change was complete in an average of 22 days after male removal, with the earliest completion of sex change happening in eight days (Warner & Swearer, 1991). However, sex reversal in T. duperrey takes approximately 60 days (Ross, 1983) with some animals completing sex reversal in as few as 49 days (Morrey et al., 2002). In undertaking this work (Larson et al. 1999), two questions were addressed. (1) Does T. duperrey show behavioral sex reversal prior to gonadal reversal, similar to other labrids? (2) Is behavioral reversal, like gonadal reversal, slower in T. duperrey than in T. bifasciatum? It was hypothesized that behavioral sex reversal would take longer in T. duperrey than in T. bifasciatum because (1) gonadal sex reversal takes almost three times longer in T. duperrey and (2) T. bifasciatum has only one male per reef whereas T. duperrey has multiple males. In order to test these hypotheses, large external tanks with flow-through seawater and small coral heads (Porites sp.) were set up to provide habitat for the fish. Fish were put into the tanks to acclimate for one week. The following populations were replicated in each of five tanks: one TP male between 140 and 150 mm standard length (SL), one large female between 120 and 130 mm SL, and four small females between 90 and 100 mm SL. Every day, T. duperrey spawn during a two-hour period around the daytime high tide. This time was selected for observations because this is when the animals show the most social interactions, especially courting and aggression (Ross, 1983). It was easy to distinguish between the TP male, the large females, and the small females because of size differences. It was not possible, however, to differentiate among the small females and they were considered as a group. After three days of observations (days 3, 2, and 1), the TP male in each tank was netted and removed. Observations of the remaining fish continued for seven days (days 1e7) following male removal. Courtship behavior consisted of looping, circling, and fluttering (Robertson & Hoffman, 1977). Looping involved swimming in an inverted U above the female. Circling

Chapter | 8


consisted of swimming in a tight circle around the female. Pectoral fluttering and quivering, sometimes performed simultaneously, consisted of rapid vibration of pectoral fins and body, respectively, in front of or above the female. In the field, spawning rushes are observed, whereby a TP male and a female swim in tandem toward the surface, releasing gametes at the apex of the rush. No spawning rushes were observed in the tanks, presumably due to space restrictions. Aggressive behaviors consisted of chasing and biting (Godwin, 1993). Prior to male removal, both the TP male and the large female (Figure 8.4) demonstrated aggression. The aggression of the TP male was mainly directed toward the large female, whereas the aggression of the large female was directed toward the smaller females. Initially, TP males and large females both demonstrated intermediate levels of aggression. Following TP male removal, aggression sharply increased in large females. Beginning with day

FIGURE 8.4 Aggressive (upper) and courting (lower) behavior in Thallasoma duperrey before and after removal of the terminal phase (TP) male (day zero). Data are means and error bars represent standard errors.

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three, this aggression tapered off and, by day five, levels were not significantly different from those prior to TP male removal. In the small females, there were no aggressive acts prior to TP male removal. Following TP male removal, aggression increased in small females and peaked on day two. After day three, aggression tapered off in the small females as well. Prior to their removal, courting behavior was exhibited only by TP males (Figure 8.4). After TP male removal, there was no courting by the large females until day three. On day three, large females began courting smaller females. This increased steadily until day seven, when courting values of large females were similar to those of TP males prior to their removal. Small females never exhibited courting behavior. Thus, prior to male removal, large females and males exhibited different behavior patterns. After male removal, the largest female underwent behavioral sex reversal and exhibited male behavior patterns. Within the genus Thalassoma, social control of sex reversal occurs in T. bifasciatum (Warner & Swearer, 1991) and Thallosoma lucasanum (Warner, 1982) as well as in T. duperrey (Ross et al., 1983; 1990). In T. duperrey, it also has been demonstrated that the relative size of individuals within the population are recognized via visual mechanisms and used as a cue. Females showed a marked increase in aggression on the first day following TP male removal. Males were removed the previous afternoon following that day’s behavioral observations. This means that the large females were not observed until 20 hours post-removal. Therefore, it is possible that aggressive behaviors could have been on the rise during the first 20 hours. The average number of aggressive attacks doubled following TP male removal. The increase in aggression exhibited by the large females subsequent to TP male removal is a common feature in most sex-changing fish (Godwin, 1993). This occurs in both protogynous (Robertson, 1972; Warner & Swearer, 1991) and protandrous (Godwin, 1993) species. Why did large females show greater aggression than TP males on day 3? On the reefs, aggression by TP males seems to be directed primarily at other TP males (Ross et al., 1983; Warner & Schultz, 1992); since there was only one TP male per tank in this experiment, the usual target of that aggression was absent. In addition, TP males in the experimental tanks were heavily involved in courting, and as such exhibited more courting than aggression (Figure 8.4). Beginning on day three, the large females showed a decrease in aggression with a concomitant increase in male courting behavior. On day four, aggression began to level out (Figure 8.4). Changes in aggressive behavior in the small females paralleled those in the large females. On day one, small females increased aggressive behavior, which was maximal

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on day two and leveled off subsequently, as did aggression in large females. Smaller females exhibited levels of aggression at roughly 45% of the large females. As there were four small females of roughly equal size in each tank, small females could not be differentiated and were treated as one group. However, most of the aggressive acts by smaller females appeared to emanate from one female. In the genus Thalassoma, relative size conveys dominance (Ross et al., 1983; Warner & Schultz, 1992) and, prior to TP male removal, the largest female was dominant and therefore the most aggressive. Following TP male removal, the position of the most dominant female was likely passed on to the most dominant of the smaller females. This was not necessarily due to size, as the females in this group did not differ by more than several millimeters, which may have resulted in elevated aggression in all of the small females as they began to establish a new social hierarchy. The most dominant of the smaller females was likely the most aggressive. Size is considered to be important in the initiation of sex reversal in T. duperrey (Ross, 1983), but it does not seem to be the ultimate factor in determining sexual or behavioral changes (Morrey et al., 2002). If a large female is placed with a TP male that is significantly smaller than she is, she will not change sex (Morrey et al., 2002). Therefore, it seems that there are factors other than just size at play in the social system of T. duperrey. As a result of all this, it is likely that the aggressive acts of the small females were attributable to a single, most dominant female establishing her position as the next heir apparent. Courting behavior in females began on day three postremoval and increased steadily until day seven, when these levels were not significantly different from those exhibited by TP males prior to removal. It is with these behaviors that the female completes the behavioral transition to TP male, and it is important to note that the large females did not exhibit courting behavior prior to TP male removal and small females never exhibited courting behavior. The decrease in aggression observed in large females on day three post-removal was likely a consequence of an increase in courting. If one looks at the behavior of the males prior to removal, aggression was at levels even lower than that of the large females and courting was high. Courting and aggression are inversely correlated among TP males and transitional animals. Behavioral reversal in T. duperrey preceded gonadal reversal. Aggression began on day 1 following male removal and courting began on day three. These behaviors may be a mechanism to prevent too many of the females from changing sex. The largest female asserts her role as the new male via these male behaviors. Her increase in aggression coupled with courting serves to keep the other females in their current positions. The timing of behavioral sex reversal in T. duperrey is slower than in T. bifasciatum. This corresponds to both the longer time course of gonadal


reversal in T. duperrey and the greater number of TP males and nonharemic mating system of this species in contrast to T. bifasciatum. Three experiments have elucidated the role of monoamines in the initiation and completion of gonadal sex reversal in T. duperrey (Larson et al., 2003b). Experiments one and two tested whether pharmacologically increasing NE or blocking DA could trigger the initiation or completion, respectively, of gonadal sex reversal in T. duperry under nonpermissive social conditions. For experiments one and two, females were housed singly, a condition that had been shown to prevent sex reversal (Ross, 1983). Time-release implants were implanted intraperitoneally. They contained drugs that either increased NE release or decreased DA or 5-HT release (Larson et al., 2003b). Experiment three tested whether adrenergic blockade or increase in DA or 5-HT could block sex reversal under permissive social conditions. This experiment tested whether monoamines are needed for the completion of sex reversal. A state of permissive social conditionsdi.e., the presence of a smaller female and the absence of a TP maledis required for T. duperrey to change sex (Ross et al., 1983). Nonpermissive social conditions are any situation in which these two criteria are not met. For the purpose of these experiments, nonpermissive social conditions refer to female isolation. Gonads were classified microscopically into one of six stages defined for socially stimulated sex reversal in T. duperrey (Nakamura et al., 1989): stage 1, a normal vitellogenic ovary; stage 2, the first step in ovarian degeneration characterized by degeneration of vitellogenic oocytes; stage 3, characterized by degeneration of previtellogenic oocytes; stage 4, the first step in becoming male, characterized by the proliferation of Leydig cells and future spermatogonia; stage 5, characterized by the onset of spermatogenesis; and stage 6, characterized by presence of sperm. As described below, all pharmacological manipulations of monoamines had an effect on the process of sex reversal. Monoaminergic signaling is both sufficient and necessary in the process of sex reversal in T. duperrey. Noradrenergic signaling appears to be an important stimulator of both the initiation and completion of the process. Serotonergic and dopaminergic mechanisms play inhibitory roles in the initiation of the process. Increasing NE via ephedrine treatment brought one animal to completion of sex reversal, while the other five all initiated testicular development (Larson et al., 2003b). Norepinephrine has been shown to stimulate the HPG axis in gonochoristic fishes (Chang & Peter, 1983b) via a adrenergic receptors (Chang et al., 1991; Yu & Peter, 1992). Norepinephrine caused an increase in circulating luteinizing hormone (LH) levels in both catfish (Heteropneustes fossilis) and trout (Oncorhynchus mykiss)

Chapter | 8


(Senthilkumaran & Joy, 1996; Linard, Bennani, & Saligaut, 1995). Koulish and Kramer (1989) used human chorionic gonadotropin (hCG) to induce sex reversal in T. bifasciatum. The action of hCG is most similar to LH in fishes. Norepinephrine acts on the HPG axis by stimulating the production of GnRH at the hypothalamus and its release to the pituitary (Chang et al., 1991). The ability of NE to cause release of GnRH in gonochoristic fishes ties in nicely with the work on GnRH in sex-changing fishes. Gonadotropinreleasing hormone-ir cell number increases with protogynous sex reversal in T. bifasciatum (Grober & Bass, 1991). Although GnRH induced sex reversal in the same species, it was in conjunction with a DA blockade (Kramer et al., 1993). Another possibility is that NE acts directly on the pituitary to cause GTH release. Multiple studies have shown adrenergic innervation of the pituitary itself. In the European eel (Anguilla anguilla), noradrenergic fibers connect the POA and the pars distalis (Fremberg, Van Veen, & Hartwig, 1977). Further, the mullet Mugil planatus has adrenergic fibers entering the rostral pars distalis (Zambrano, 1975). In T. duperrey, increasing synaptic NE was effective in both the initiation and completion of sex reversal. Blocking NE action by blocking a-receptors effectively prevented the animals from progressing through the male phases of sex reversal. Blocking DA receptors with haloperidol initiated sex reversal in T. duperrey. Conversely, APO prevented animals from completing sex reversal under permissive social conditions (Figure 8.5). The effect of DA on sex reversal, as with NE, may be a function of its effects on GnRH or via direct pituitary innervation. Several investigators have shown direct innervation of the pituitary by dopaminergic neurons. Dopaminergic perikarya located in the preoptic recess organ project to the pars distalis (Kah, Dulka, Dubourg, Thibault, & Peter, 1987). Such dopaminergic neurons likely are involved in the neuroendocrine regulation of the anterior pituitary functions, in particular the inhibition of GTH release (Peter et al., 1986). The results of this study were very compelling, especially when compared with previous results concerning the physiology of protogyny. Kramer et al. (1993) only successfully stimulated sex reversal in T. bifasciatum when GnRH was combined with a DA antagonist. However, haloperidol alone was successful in the formation of spermatogenetic crypts in two of four T. duperrey (Figure 8.5). This suggests that inhibition of DA is more important for the initiation of sex reversal, at least in T. duperrey, but likely in other species as well. Although GnRH is likely an important component in the regulation of sex reversal, the evidence that it cannot initiate sex reversal without dopaminergic inhibition and that dopaminergic inhibition alone is sufficient suggests that GnRH is more important for reactivation of gonadal function after the animal becomes a male. Therefore, in T. duperrey and likely in

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FIGURE 8.5 Pharmacological manipulation of monoamine neurotransmitters in Thalassoma duperrey. (a) Stimulation under nonpermissive conditions. Ephedrine and maprotiline stimulate the noradrenergic system, while haldol inhibits the dopaminergic system. (b) Inhibition under permissive conditions. Phenoxybenzamine (PBZ) inhibits the noradrenergic system, while apomorphine (APO) and sertraline stimulate the dopaminergic and serotonergic systems respectively. Data are means and error bars represent standard errors. Bars without SEM indicate that all animals were at the same stage. Bars marked with the same letter were not significantly different from each other. Reproduced from Larson, Norris, and Summers (2003).

T. bifasciatum as well, DA seems very important in the initiation of sex reversal. Blocking DA appeared to be the most effective means of initiating sex reversal. Dopamine blockade, however, did not promote completion of sex reversal. Dopaminergic stimulation did prevent the completion of sex reversal (stage 6), but it did not keep fish from entering the early male stages of gonadal sex reversal (stages 4 or 5). Serotonergic manipulations were also effective in altering the process of sex reversal. Animals treated with

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the 5-HT antagonist ritanserin under nonpermissive social conditions (Figure 8.5) underwent sex reversal. Fish treated with the 5-HT reuptake inhibitor sertraline did not change sex under permissive social conditions. Increasing synaptic 5-HT seemed the most effective method for arresting the process at an early stage. Sex reversal under natural conditions is a social event, and the effects of social stress are likely to be mediated by 5-HT. A role for 5-HT in social interactions, dominance hierarchies, and aggression is well-substantiated in fishes (Winberg & Nilsson, 1993). Serotonin production and metabolism are inversely related to position in the dominance hierarchy for Arctic char (S. alpinus) (Winberg et al., 1991). Perciform fishes such as the cichlid A. burtoni and the damselfish Pomacentrus partitus, two species in the same suborder as T. duperrey, have serotonergic components of the dominance hierarchies. In the damselfish, socially interacting individuals had higher 5-HT than did isolated fish, with subordinate individuals having higher 5-HT metabolism than did dominant males (Winberg, Myrberg, & Nilsson, 1997). In A. burtoni, subordinate males had greater 5-HT metabolism than did dominant males (S. Winberg, Y. Winberg, & Fernald, 1996). Sexual maturation is suppressed in the subordinate males of this species, unlike the case for wrasses. Suppression of sexual maturation via serotonergic mechanisms suggests a possible mechanism for the serotonergic control of sex reversal. Serotonin modulates both NE and DA. For example, in the rat hypothalamus, stimulation of 5-HT1 and 5-HT2 receptors causes a decrease in NE release, whereas 5-HT3 stimulation results in a decrease in NE release (Blandina, Goldfarb, Walcott, & Green, 1991). Perhaps monoamine systems are integrated during activation of behavioral and gonadal events during sex reversal in teleosts. While this has not been previously demonstrated in sex-changing fishes, there have been studies in other fishes that suggest that the 5-HT system could be tied to reproduction similarly to the way it is in mammals. In the Atlantic croaker (Micropogonias undulatus), 5-HT immunoreactivity shows a similar neuroanatomical pattern to that in mammals and shares a similar proximity with the GnRH system (Khan & Thomas, 1993). This supports the role of 5HT in reproduction in at least one teleost species. African catfish (Clarias gariepinus) and Senegalese sole (Solea senegalensis) show similar serotonergic neuroanatomy (Corio, Peute, & Steinbusch, 1991; Rodrı´guez-Go´mez, Rendoń-Unceta, Sarasquete, & Munõz-Cueto, 2000). Even sharks show similarities with mammals in the development and anatomy of the 5-HT system (Carrera, Molist, Anado´, & Rodrı´guez-Moldes, 2008). In addition to internal gonadal changes, many studies have looked for external color changes as evidence of sex reversal. Although T. duperrey does not have the degree of dichromatism exhibited by other labrids, there are subtle color differences between IP and TP males. With


T. duperrey, many of the pharmacologically treated animals exhibited the external coloration and morphological characteristics of TP males in addition to undergoing gonadal changes (Larson et al., 2003b). Terminal color phase was observed in eight out of ten individuals with increased NE, but not in any of the other treatments. There are two possible explanations for NE associated with color change. As NE is the most effective stimulator of the HPG axis, it may be causing color change via stimulation of androgen production. The other explanation is that NE is driving color change via direct action on chromatophores, as demonstrated in various species of fishes (Kasukawa, Sugimoto, Oshima, & Fujii, 1985). These experiments demonstrate that monoamines are important contributors in the control of sex reversal (Larson et al., 2003b). Norepinephrine is an important stimulator for the initiation and completion of gonadal sex reversal as well as color change. Dopamine exercises inhibitory action on the initiation of sex reversal while 5-HT inhibits sex reversal most likely through its effects on NE. Thus, the serotonergic system appears to be an integral part of the pathway mediating the conversion of a social cue into a neuroendocrine event. Now that the timing of sex reversal had been established and it had been shown that pharmacological manipulations of monoamines can affect sex reversal (Larson et al., 2003b), it was time to test the finial aspect by measuring monoamine levels throughout sex reversal. Previous work on the physiology of sex reversal coupled with the in-vivo pharmacological manipulation of monoamines in T. duperrey (Larson et al., 2003b) led to the following hypothesis: sex reversal is accompanied by an increase in noradrenergic activity along with a decrease in both serotonergic and dopaminergic activity. Two females were housed together to induce sex reversal in the larger female (Ross et al., 1983), which was removed at various time points in the process of sex reversal for monoamine analysis (Larson et al., 2003a). Monoamine turnover was estimated using the ratio of metabolite to parent amine (e.g., 3-methoxy-4-hydroxy phenylglycol (MHPG): NE). This ratio is used frequently as an indirect measure of the rate of catabolism for the transmitter and indicates transmitter turnover and system activity. The most dramatic changes in monoamine turnover were seen during the first week of sex reversal (Larson et al., 2003a). It is during this time that transitional animals undergo behavioral sex reversal (Larson, Norris, Grau, & Summers, 1999). In the POA, there were marked changes in the noradrenergic and serotonergic systems (Larson et al., 2003a). Noradrenergic activity started at intermediate levels on day one and drastically increased at day three (Figure 8.6). Beginning with day five, levels returned to their previous state (Figure 8.6). Serotonergic activity was moderate on day one, dropping sharply on day three (Figure 8.6). On

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FIGURE 8.6 Changes in noradrenergic (3-methoxy-4-hydroxy phenylglycol (MHPG) : norepinephrine (NE)) and serotonergic (5-hydroxy indoleacetic acid (HIAA): serotonin (5-HT)) activities in Thallasoma duperrey during the first week of male-to-female sex reversal, as indicated by metabolite to transmitter ratios. Day one represents one day after male removal. The upper panel represents the preoptic area (POA) and the lower represents the raphe nucleus (RN). Data are means and error bars represent standard errors.

days five and seven, there was an increase in turnover (Figure 8.6). The raphe nucleus (RN) exhibited marked changes in serotonergic activity (Larson et al., 2003a). Serotonergic activity was high on day one, dropping on days three (2.0058, n ¼ 3) and five and then increasing again on day seven (Figure 8.6). Monoamines appear to play a significant role in regulating sex reversal. Changes were demonstrated in monoamine metabolism for all brain regions examined, albeit not for every neurotransmitter system in each region. The most important changes in monoamine system activation were seen during the first week of sex reversal, particularly in the

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POA and the RN. It is during this time that transitional animals undergo behavioral sex reversal. Noradrenergic activity in the POA, a region of the brain that traditionally is associated with male sexual behavior and function in vertebrates, of T. duperrey was low on day one, drastically increased on day three, and returned to the previous level of activity on day five (Figure 8.6), suggesting a brief period of noradrenergic activity that perhaps triggers courting on day three. Sex-changing females began to exhibit courting on day three following removal of the TP male (Larson et al., 2003a). Additionally, the observed increase in NE metabolism with simultaneous changes in 5-HT metabolism may have implications with respect to activity of the HPG axis. Serotonin activity decreased on days three and five and then rose again on day seven. Serotonin is highly responsive to social stress in vertebrates including fishes (Winberg & Nilsson, 1993). Serotonin inhibits the noradrenergic activity in the POA of rats (Blandina et al., 1991), where NE stimulates the HPG axis (Peter et al., 1986). A picture emerges for T. duperrey in which serotonergic activity decreases in response to male removal, allowing noradrenergic activity to peak, and this sends a signal along the HPG axis, triggering gonadal sex change. The RN contains large numbers of neuronal cell bodies producing 5-HT that project to virtually every other part of the vertebrate brain (Butler & Hodos, 1996). On days one through five there is a sharp decrease in serotonergic activity in the locus coeruleus (LC). It is during this time that it is most crucial for the female to assert dominance over the other females. Serotonin plays a very important role in the acquisition of dominance (Larson & Summers, 2001), especially in the RN (Kostowski, Plewako, & Bidzinski, 1984). In these systems, low serotonergic activity conveys dominance and pharmacologically enhancing serotonergic activity can reverse dominance in both reptiles and mammals (Kostowski et al., 1984; Larson & Summers, 2001). This decrease in the serotonergic activity in the RN of T. duperrey between days one and five may be very important for the female who was previously second in the dominance hierarchy and is now the alpha (most dominant) individual. Additionally, serotonergic activity in the RN can play an inhibitory role in male sexual behavior. In male rats, injection of the 5-HT synthesis inhibitor p-chlorophenylalanine (PCPA) into the RN increases the amount of male sexual behavior (Yamanouchi & Kakeyama, 1992). Injecting PCPA into the RN of female rats also induces male sexual behavior (Matsumoto & Yamanouchi, 1997). Results of PCPA in the mammalian RN have implications for sex change in T. duperrey. By injecting the PCPA into the RN of rats, the pharmacologically induced decrease in serotonergic activity causes an increase in male behavior in both male and female rats (Yamanouchi & Kakeyama,

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1992; Matsumoto & Yamanouchi, 1997). In T. duperrey, females exhibit an increase in male sexual behavior during the transition from female to male. Concomitant with the change from female to male sexual behavior is a decrease in serotonergic activity in the RN. This suggests that the decrease in serotonin in the RN on day three is linked to, and perhaps causes, the behavioral sex reversal. The study of Larson et al. (2003a) was the first to investigate monoamines in relation to natural sex reversal and revealed changes in monoaminergic activity in all brain regions investigated. The most striking changes occurred in the POA and the RN. In the POA, a vast reorganization of all monoaminergic systems occurs on day three, the day that courting is first observed. There is an increase in noradrenergic activity concomitant with drastic decreases in both serotonergic and dopaminergic activity. The likely explanation is that the decrease in serotonergic activity due to male removal produces the increase in noradrenergic activity, which, coupled with the decrease in dopaminergic activity, triggers gonadal sex reversal. There was a drastic decrease in the serotonergic activity in the RN from day one to five that may be important both in the acquisition of dominance and the transition from female to male sexual behaviors. Since this study, other work has looked at the role of 5-HT in sex reversal. Perrault, Semsar, and Godwin (2003) showed that increasing 5-HT with the 5-HT reuptake inhibitor fluoxetine decreases aggression in TP males of T. bifasciatum. This supports the role of 5-HT in decreasing dominance and aggression. Fluoxetinetreated TP males also show a decrease in AVT expression in the POA (Semsar, Perrault, and Godwin, 2004). These results tie in well with a decrease in aggression with decreasing AVT and support not only the idea that both 5-HT and AVT are important in sex-changing fishes but also that there are many different components that play a role. The rapid change in expression of the monoamines during sex reversal is an important step in the complex reorganization of the entire HPG axis during this process, involving not only monoamines but also steroids and various peptides.

7. FUTURE RESEARCH Future research into the neuroendocrine control of sex change in fishes offers many possibilities. Sex reversal is a very complex psychosocial and physiological phenomenon. There are many unanswered questions about the different factors responsible for behavioral vs. gonadal sex reversal. Recently the field of genomics has undergone a dramatic expansion. While there have yet to be any studies on the genome of any sex-changing fish, it is only a matter of time. Other fish have genome projects, e.g. the zebrafish, fugu, and salmon (Elgar et al., 1996; Postlethwait, Amores, Force, Yan, and Billard, 1999;


Adzhubei, 2007). Additionally, microarray work has been done on other species such as cichlids and swordtails, looking at the relationship between genomics and social or sexual behavior (Cummings et al. 2008; Renn, AubinHorth, and Hofmann, 2008). These types of studies should be done with sex-changing species. It has yet to be seen what possibilities and challenges genomics may hold for the study of sex-changing fishes, but perhaps the technology may be capable of answering some of the unanswered questions. Additionally, continuing to link physiology with behavior in more traditional ways still offers many possibilities for filling gaps in our knowledge. Understanding the physiological and behavioral processes that interact to orchestrate sex change in fishes is a complicated puzzle in which we still have many pieces to place.

ABBREVIATIONS 11-KT 5-HT AND APO AVT BSD DA DHEA E2 ESD GnRH GTH hCG HPG IP ir LC LH MHPG NE PCPA POA RN RV SAH SL T TP

11-ketotestosterone Serotonin Androstenedione Apomorphine Arginine vasotocin Behavioral sex determination Dopamine Dehydroepiandrosterone Estradiol Environmental sex determination Gonadotropin-releasing hormone Gonadotropin Human chorionic gonadotropin Hypothalamicepituitaryegonadal Initial phase Immunoreactive Locus coeruleus Luteinizing hormone 3-methoxy-4-hydroxy phenylglycol Norepinephrine P-chlorophenylalanine Preoptic area Raphe nucleus Reproductive value Size advantage hypothesis Standard length Testosterone Terminal phase

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Shapiro, D. Y. (1981). Behavioural changes of protogynous sex reversal in a coral reef fish in the laboratory. Anim. Behav., 29, 1185e1198. Shapiro, D. Y. (1988). Behavioral influences on gene structure and other new ideas concerning sex change in fishes. Environ. Biol. Fish., 23, 283e297. Shapiro, D. Y. (1992). Plasticity of gonadal development and protandry in fishes. J. Exp. Zool., 261, 194e203. Shapiro, D. Y., & Sadovy, Y. (1993). Periodicity of sex change and reproduction in the red hind, epinephelus guttatus, a protogynous grouper. B. Mar. Sci., 53, 1151e1162. Shapiro, D. Y., Marconato, A., & Yoshikawa, T. (1994). Sperm economy in a coral reef fish, Thalassoma bifasciatum. Ecology., 75, 194e203. Shen, S.-C., & Liu, C. H. (1976). Ecological and morphological study of the fish fauna from the waters around Taiwan and its adjacent islands. Acta Oceanogr. Taiwan, 6, 140e150. Shinomiya, A., Yamada, M., & Sunobe, T. (2003). Mating system and protandrous sex change in the lizard flathead, Inegocia japonica (Platycephalidae). Ichthyol. Res., 50, 383e386. Smith, C. L. (1982). Patterns of reproduction in coral reef fishes. In G. R. Huntsman, W. R. Nicholson, & W. W. Fox (Eds.), The biological bases for reef fishery management; NOAA Tech. Memo. NMFS-SEFC-80 (pp. 49e66). Washington, DC: NOAA. Sneddon, L. U., Margareto, J., & Cossins, A. R. (2005). The use of transcriptomics to address questions in behaviour: production of a suppression subtractive hybridisation library from dominance hierarchies of rainbow trout. Physiol. Biochem. Zool., 78, 695e705. Somoza, G. M., Yu, K. L., & Peter, R. E. (1988). Serotonin stimulates gonadotropin release in female and male goldfish, Carassius auratus L. Gen. Comp. Endocrinol., 72, 374e382. St. Mary, C. M. (1993). Novel sexual patterns in two simultaneous hermaphroditic gobies Lythrypnus dalli and Lythrypnua zebra. Copeia, 4, 1062e1072. Stoll, L. M. (1955). Hormonal control of the sexually dimorphic pigmentation of Thalassoma bifasciatum. Zoologica, 40, 125e132. Streelman, J. T., Alfaro, M., Weastneat, M. W., Bellwood, D. R., & Karl, S. A. (2002). Evolutionary history of the parrotfishes: Biogeography, ecomorphology, and comparative diversity. Evolution, 56, 961e971. Sunobe, T., & Nakazono, A. (1993). Sex change in both directions by alteration of social dominance in Trimma okinawae (Pisces: Gobiidae). Ethology, 94, 339e345. Sunobe, T., Nakamura, M., Kobayashi, Y., Kobayashi, T., & Nagahama, Y. (2005). Aromatase immunoreactivity and the role of enzymes in steroid pathways for inducing sex change in the hermaphrodite gobiid fish Trimma okinawae. Comp. Biochem. Physiol. A. Mol. Integr. Physiol., 141, 54e59. Tchoudakova, A., & Callard, G. V. (1998). Identification of multiple cyp19 genes encoding different cytochrome P450 aromatase isozymes in brain and ovary. Endocrinology, 139, 2179e2189. Tan-Fermin, J. D. (1992). Withdrawal of exogenous 17-alpha methyltestosterone causes reversal of sex-inversed male grouper Epinephelus suillus (Bloch and Schneider). Philippine Sci., 29, 33e39. Trant, J. M., Gavasso, S., Ackers, J., Chung, B. C., & Place, A. R. (2001). Developmental expression of cytochrome P450 aromatase genes (cyp19a and cyp19b) in zebrafish fry (Danio rerio). J. Exp. Zool., 290, 475e483.


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Chapter 9

Hormonally Derived Sex Pheromones in Fishes Norm Stacey University of Alberta, Edmonton, AB, Canada

SUMMARY Fishes commonly use reproductive hormones and their precursors and metabolites as ‘hormonal pheromones’ that induce important behavioral and physiological effects in conspecifics. In goldfish, the best understood example, periovulatory females release steroid and prostaglandin pheromones that increase male hormone levels, sexual behaviors, ejaculate volume, sperm motility, and paternity. Although such reproductive responses have been described in only a dozen species, olfactory recordings show that several hundred species from major orders (Cypriniformes, Characiformes, Siluriformes, Salmoniformes, Perciformes) are extraordinarily sensitive (picomolar threshold) to water-borne hormonal compounds. Moreover, a number of these species discriminate a variety of hormonal compounds through multiple, highly specific olfactory receptor mechanisms, suggesting that, despite the limited chemical diversity of hormones per se, hormonal pheromones might be species-specific. The discovery of hormonal pheromones broadens the classical concept of hormone actions being restricted to the individual by showing that many aspects of fish reproductive function result from exogenous hormonal effects among conspecifics.

1. INTRODUCTION Vertebrate reproductive hormones promote sexual synchrony among conspecifics both by acting on the brain to synchronize individual reproductive behavior with the appropriate stage of gamete maturation, and by acting on effectors to generate signals that synchronize reproduction between individuals. In fishes, however, hormonal steroids and prostaglandins that are released into the water can promote reproductive synchrony through an important third action, by acting as potent pheromones that induce physiological and behavioral responses in conspecifics (Stacey & Sorensen, 2006; Stacey, 2009; Stacey & Sorensen, 2009). In this chapter, such pheromones will be referred to as ‘hormonal pheromones,’ regardless of whether they consist only of hormones per se, only of hormonal


precursors and metabolites, or both. Although not well understood, hormonal pheromones are likely to be multicomponent; e.g., the preovulatory steroid pheromone of goldfish (Carassius auratus), which evidently consists of at least three steroids (Kobayashi, Sorensen, & Stacey, 2002). Implicit in the term ‘hormonal pheromone’ is the assumption that, because reproductive events are expected to be preceded not only by increased release of hormones but also by increased release of hormonal precursors and metabolites, it is unnecessary to distinguish between pheromones consisting of hormones and pheromones consisting of hormonal proxies. Although it has long been known that fishes have commonly evolved reproductive pheromones with diverse physiological (‘primer’) and behavioral (‘releaser’) effects on conspecifics (Liley, 1982; Burnard, Gozlan, & Griffiths, 2008), only recently has it become clear that many reproductive pheromones of fishes might be comprised solely or partially of hormones and related compounds from hormone synthesis pathways. Døving (1976) appears to have been the first to propose that, living in a medium where visual cues are often limited but where the olfactory organ is readily exposed to soluble conspecific odors, fishes are predisposed to evolve pheromonal functions for released sex hormones, because these informationrich chemicals should be reliable indicators of key reproductive events. In support, Colombo, Marconato, Belvedere, and Frisco (1980) soon reported that a conjugated testicular steroid (5b-androstan-3a,17b-diol-3aglucuronide ¼ etiocholanolone glucuronide or Etio-g) induced pheromonal effects in the black goby (Gobius niger), and Van den Hurk and Lambert (1983) (see also Van den Hurk & Resink, 1992) reported that glucuronides of testosterone (T-g) and 17b-estradiol (E2-3g) attracted male zebrafish (Danio rerio). In the following 25 years, steroids, prostaglandins, and their precursors and metabolites have been found to have primer and releaser effects in a variety of fishes from four major orders: Cypriniformes (goldfish 169

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(reviewed by Kobayashi et al., 2002; Stacey & Sorensen, 2006; 2009); crucian carp (Carassius carassius) (Bjerselius, Olseń, & Zheng, 1995; Olseń, Sawisky, & Stacey, 2006); common carp (Cyprinus carpio) (Irvine & Sorensen, 1993; Stacey, Zheng, & Cardwell, 1994); oriental weatherfish loach (Misgurnus anguillicaudatus) (Kitamura, Ogata, & Takashima, 1994a); tinfoil barbs (Barbonymus spp.) (Cardwell, Stacey, Tan, McAdam, & Lang, 1995)); Siluriformes (African catfish (Clarias gariepinus) (reviewed by Van den Hurk & Resink, 1992)); Salmoniformes (Atlantic salmon (Salmo salar) (Moore & Waring, 1996); brown trout (Salmo trutta) (Moore, Olseń, Lower, & Kindahl, 2002); lake whitefish (Coregonus clupeaformis) (Laberge & Hara, 2003); arctic char (Salvelinus alpinus) (Sveinsson & Hara, 2000)), and Perciformes (round goby (Neogobius melanostomus) (Murphy, Stacey, & Corkum, 2001)). In addition to these reports that water-borne hormones induce physiological and behavioral effects in over a dozen fish species, studies using underwater electro-olfactogram (EOG) recording (Scott & Scott-Johnson, 2002) to screen for olfactory responsiveness to a large number of prostaglandins and steroids have demonstrated that well over a hundred additional species are acutely sensitive to these compounds. Thus, in the primarily freshwater superorder Ostariophysi (Nelson, 2006), virtually all of more than 100 cypriniform (carps and minnows), siluriform (catfish) (Narayanan & Stacey, 2003), and characiform fishes (Cardwell & Stacey, 1995) detected at least one prostaglandin or steroid at the low (pM) concentrations expected of a pheromone derived from hormonal products that already are at extremely low (nM) concentrations in plasma prior to release (Stacey & Sorensen, 2006; 2009). Moreover, such olfactory responsiveness to hormones and synthetically related compounds occurs across a broad array of fishes, from primitive Elopiformes (Megalops cyprinoides (tarpons) (Stacey & Sorensen, 2006; 2009)) to highly derived Perciformes (Gobiidae (Murphy et al., 2001); Cichlidae (Cole & Stacey, 2006)), suggesting that it is reasonable to expect that hormonal pheromones are widely distributed among fishes. Indeed, given the extraordinary specificity of the olfactory receptor mechanisms detecting steroids and prostaglandins (Sorensen, Hara, Stacey, & Goetz, 1988; Sorensen, Hara, Stacey, & Dulka, 1990) and the fact that many hormonal pheromones may be unknown hormonal metabolites (e.g., Barata et al., 2008), it is expected that these EOG screening studies greatly underestimate both the range of hormonal compounds detected by fishes, and the number of fish that detect them. Although hormonal pheromones are not well understood in any fish, it already is clear from studies of goldfish, arguably the most informative example, that knowledge of a species’ reproduction might be very incomplete if nothing


is known of its hormonal pheromones. More generally, there are two important ways in which the discovery of hormonal pheromones broadens our perspective of fish reproduction. The first is the simple demonstration that actions of the reproductive endocrine system are not restricted to endogenous hormonal control, but commonly encompass equally important exogenous pheromonal effects (Stacey, Chojnacki, Narayanan, Cole, & Murphy, 2003). The second is that, where such pheromones induce hormonal change, there is the possibility that this response of receivers might not only generate feedback to the pheromone releaser but also relay the pheromonal signal to additional conspecifics. In such situations, where hormonal pheromones act as exogenous links among the endocrine systems of numerous conspecifics (and the goldfish appears to be an example (Kobayashi et al., 2002; Wisenden & Stacey, 2005)), it might in fact be appropriate to speak in terms of ectohormones synchronizing a metaendocrine system. Ironically, Bethe (1932) proposed the term ‘ectohormone’ to describe chemicals functioning in intraspecific communication, whereas Karlson and Lu¨scher (1959) sought to restore a clear distinction between chemically mediated endogenous and exogenous functions by proposing that ‘ectohormone’ be replaced by the new term ‘pheromone’. Perhaps the most intriguing issue arising from the discovery of fish hormonal pheromones is that, being derived from a homologous and highly conserved vertebrate endocrine system, they might be considered incapable of conveying the species-specific information often considered a hallmark of pheromone function. Although this very important question has not yet been directly addressed, EOG studies indicate that congeners can detect very similar hormonal products (Irvine & Sorensen, 1993; Narayanan & Stacey, 2003) suggesting that, if it occurs, specificity likely is achieved by altering the ratios of common mixtures, or by addition of nonhormonal compounds. Electro-olfactogram studies also show that, unlike the situation among lower taxa, patterns of detected compounds can differ dramatically among higher taxa and, as discussed below (see Section 2.3.3), appear to provide examples of multiple pheromonal origins and subsequent divergence that point to numerous opportunities for comparative studies of pheromone function and evolution (Sorensen & Stacey, 1999; Stacey & Sorensen, 2006; 2009). As a result of the particularly strong correspondence between hormonal pheromones and phylogeny, this brief review summarizes current information on hormonal pheromones from a phylogenetic perspective.

2. HORMONAL PHEROMONES IN FISHES The term ‘fish’ refers to members of a paraphyletic group of more than 28 000 vertebrates that excludes tetrapods, has

Chapter | 9

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Hormonally Derived Sex Pheromones in Fishes

limbs in the shape of fins, and uses gills for aquatic respiration (Nelson, 2006). Although each of the five living classes of craniate vertebrates is comprised entirely (Myxini, Petromyzontida, Chondrichthyes, Actinopterygii) or partially (Sarcopterygii) of fishes, hormonal pheromones have been described only in the actinopterygian or ray-finned fishes, which include more than 95% of extant fishes (Nelson, 2006). In petromyzontids, however, sea lamprey (Petromyzon marinus) appear to have evolved a comparable pheromone system in which released bile steroids have been co-opted to serve as pheromones that function both as migratory attractants and sexual signals (Li et al., 2002; Sorensen & Hoye, 2007; Fine & Sorensen, 2008). Teleost fishes constitute the great majority of actinopterygians, and are the only fishes in which hormonal pheromones have been reported. Teleosts include two minor subdivisions of basal fishes (osteoglossomorphsd about 200 species of mooneyes and bonytongues; elopomorphsdabout 850 species of ten-pounders, tarpons, and eels) and two major subdivisions of more derived fishes

(the ostarioclupeomorphs or otocephalans and the euteleosts) (Table 9.1) (Nelson, 2006). Osteoclupeomorphs (about 8300 species) include clupeomorphs (herrings) and the ostariophysan cypriniforms (carps and minnows), characiforms (piranha, tetras), siluriforms (catfishes), gonorhynchiforms (milkfishes), and gymnotiforms (New World knifefishes). Despite attempts to chemically characterize a potent pheromone in milt (seminal fluid) of Pacific herring (Clupea pallasii) (e.g., Carolsfeld, Scott, & Sherwood, 1997), there is no evidence for hormonal pheromones in clupeiforms. In cypriniforms, characiforms, and siluriforms, however, hormonal pheromones evidently are widespread. Of the osteoclupeomorphs, other reviews (Van den Hurk & Resink, 1992; Sorensen & Stacey, 1999; Stacey & Sorensen, 2002; 2006) have summarized the earlier studies on zebrafish (Van den Hurk & Lambert, 1983), catfish (Resink, Voorthuis, Van den Hurk, Peters, & Van Oordt, 1989; Narayanan & Stacey, 2003), and characiforms (Cardwell & Stacey, 1995), and only goldfish and closely related cypriniforms will be discussed here. The euteleosts, sister group to the

TABLE 9.1 Electro-olfactogram (EOG) evidence for detection of prostaglandins (PGs) and steroids in orders of teleostean fishes1,2 Evidence for Hormonal Pheromones Number of extant species

Order

Number of species examined

Steroids

Common names

Example genera

PGs

Unconjugated

Conjugated

tarpon

Megalops

þ5

þ

06

Osteoglossomorph Orders Elopiformes3

8

1

Cypriniformes

3,200

>80

cyprinids

Carassius; Danio

þ

þ

þ

Characiformes

1,600

>20

characins

Astyanax; Colossoma

þ

þ

þ

Siluriformes

2,800

>20

catfishes

Clarias, Synodontis

þ

þ

þ

170

1

knifefishes

Apteronotus

þ

0

0

88

1

smelts

Plecoglossus

þ

0

0

66

9

salmonids

Salmo, Oncorhynchus

þ

þ

þ

Cyprinodontiformes

1,000

1

rivulines

Aplocheilus

þ

Perciformes

9,293

>75

cichlids, gobies

Astatotilapia, Neogobius

0

þ

þ

Osteoclupeomorph Orders

Gymnotiformes Euteleost Orders Osmeriformes4 Salmoniformes 3

1

Systematic terminology from Nelson (2006). After Stacey and Sorensen (2009), with permission, and Stacey (2009), with permission. Stacey (unpublished results). 4 See Kitamura et al. (1994b). 5 Some of the tested species in the taxon responded to at least one compound in this category. 6 No evidence that species in the taxon detect compounds in this category. 2 3

172


osteoclupeomorphs, include 28 orders with about 17 500 species: hormonal pheromones have been studied in detail in only two euteleost orders, Salmoniformes and Perciformes (gobies and cichlids) (Table 9.1).

Goldfish hormonal pheromones are better understood than those of other fishes (Kobayashi et al., 2002; Stacey & Sorensen, 2006; 2009), although undoubtedly this understanding is far from complete. Male goldfish release a hormonal pheromone (Sorensen, Pinillos, & Scott, 2005), but much more is known about females, which sequentially release three pheromones, two of which are mixtures of hormones and related compounds (Figure 9.1). Although less well studied, the hormonal pheromones of the closely related crucian and common carps appear remarkably similar to those of goldfish (Irvine & Sorensen, 1993; Stacey et al., 1994; Bjerselius et al., 1995; Olseń et al., 2006), despite the fact that goldfish and carp clearly discriminate each other’s nonreproductive whole body odors (Sisler & Sorensen, 2008). From vitellogenesis until the completion of spawning, recrudescent and periovulatory female goldfish sequentially release three pheromones that affect the behavior and physiology of males (Kobayashi et al., 2002) (Figure 9.1). The uncharacterized recrudescence pheromone, which

2.1. Goldfish and Related Cypriniforms Cypriniforms include superfamily Cyprinoidea (family Cyprinidae: about 2400 species of carps and minnows) and superfamily Cobitoidea (about 850 species) including the families Cobitidae (loaches), Catostomidae (suckers), Gyrinocheilidae (algae eaters), and Balitoridae (river loaches) (Nelson, 2006). In EOG screening studies (Stacey and Sorensen, 2006; Stacey, 2009; Stacey and Sorensen, 2009), all of more than 80 cypriniform species detected F2-series prostaglandins (PGFs), whereas detection of free and conjugated (sulfated and glucuronidated) steroids was observed only in cyprinids, in which all major taxa except acheilognathins (bitterlings) were responsive.

Oviposition OOCYTE MATURATION

VITELLOGENESIS

FEMALE

LH

E2

17,20 β -P

OVULATION behavior

17,20β -P-s

PGF2α

AND

?

Release to the Water

Preovulatory pheromone

Recrudescent pheromone

MALE

Attraction to female

AND 17,20β-P 17,20β-P-s

Agonistic behavior

LH

PGF2 α 15K-PGF2α

Postovulatory pheromone

Following + chasing Courtship + spawning behaviors

Milt

1200

SPAWNING SYNCHRONY

2000

Time of day

0400

1200

FIGURE 9.1 Schematic model of female goldfish pheromones and their physiological (primer) and behavioral (releaser) effects on males. During vitellogenesis, 17b-estradiol (E2) stimulates urinary release of an unidentified recrudescent pheromone that attracts males. In postvitellogenic females, exogenous cues induce a luteinizing hormone (LH) surge that stimulates release of a preovulatory steroid pheromone containing androstenedione (AND), the maturation-inducing steroid 17,20b-P, and its sulfated metabolite 17,20b-P-s. Early in the LH surge, AND induces agonistic behavior among males, but, as the ratio of 17,20b-P : AND increases, males increase LH and begin to follow and chase conspecifics. Late in the LH surge, 17,20b-P-s dominates the pheromone mixture, enhancing its endocrine and behavioral effects. Males exposed to the preovulatory pheromone increase both the quantity and quality of their sperm stores prior to ovulation, and increase their ejaculate volume and paternity during spawning. At ovulation, eggs evidently stimulate the oviduct to synthesize prostaglandin F2a (PGF2a), which acts in the brain to stimulate female sexual behavior and is released in urinary pulses with its major metabolite (15-keto-PGF2a) as a postovulatory pheromone stimulating both male sexual behaviors and LH release. Reprinted with permission from: Kobayashi et al. (2002). Stacey and Sorensen. 2006. Stacey and Sorensen. (2008). Stacey and Sorensen. (2009). Stacey. (2009).

Chapter | 9


attracts males and thereby might function to maintain mixed-sex aggregations, is induced by 17b-estradiol (E2) and appears to be released in urine throughout vitellogenesis (Yamazaki, 1990). In contrast, release of the preovulatory steroid pheromone and the postovulatory prostaglandin pheromone occurs only during the brief periovulatory period between the onset of the preovulatory luteinizing hormone (LH) surge and completion of oviposition. As with crucian and common carp, goldfish are springesummer spawners that have little external sexual dimorphism and live in apparently unstructured, mixed-sex groups in temperate waters. It seems likely that hormonal pheromone evolution in these carps has been driven by two major selective forces: a typically dimly lit and turbid spawning environment, and a scramble competition or polygynandrous mating system (Taborsky, 2001; Taylor & Knight, 2008) in which males engage in intense sperm competition to fertilize eggs, the sole determinant of their reproductive success. A cascade of endocrine and physiological changes that lead to spawning begins when increasing water temperature and aquatic vegetation (substrate for the adhesive eggs) trigger an afternoon neuroendocrine reflex, resulting in a photoperiodically synchronized LH surge that stimulates follicular synthesis of the oocyte maturation-inducing steroid 17,20b-dihydroxy-4-pregnen-3-one (17,20b-P) (Stacey, Cook, & Peter, 1979; Kobayashi et al., 2002; Canosa, Stacey, & Peter, 2008). When ovulation occurs, approximately 12 hours later, females become sexually active for the several hours that eggs in the oviduct stimulate synthesis of prostaglandin F2a (PGF2a), which acts within the brain to stimulate female oviposition behavior (Stacey & Peter, 1979; Sorensen et al., 1995a) (Figure 9.1). Small groups of males then vigorously compete for access to ovulated females as they repeatedly enter aquatic vegetation to oviposit large numbers of unguarded eggs over a period of several hours. Throughout the approximately 15 hours between onset of the LH surge and completion of oviposition, the female releases a preovulatory steroid pheromone and a postovulatory prostaglandin pheromone, both of which dramatically affect the physiology and behavior of males.

2.1.1. Female preovulatory steroid pheromone Preovulatory steroid pheromone release begins soon after onset of the preovulatory LH surge and terminates at ovulation the following morning (Stacey et al., 1979; Stacey, Sorensen, Van der Kraak, & Dulka, 1989; Canosa et al., 2008) (Figure 9.1). The odor of females undergoing a preovulatory LH surge induces in intact males, but not in anosmic males (Kobayashi, Aida, & Hanyu, 1986a), rapid increases in plasma LH and steroid concentrations, which

173

in turn increase the volume of milt (sperm and seminal fluid) in the sperm ducts within several hours (Kobayashi Aida, & Hanyu, 1986b; Dulka, Stacey, Sorensen, & Van der Kraak, 1987; Stacey et al., 1989). For simplicity, the odor females release during this period is referred to as a single preovulatory pheromone; however, the behavioral and physiological responses of males indicate that they perceive multiple odors that result from the pheromone’s components varying both temporally, due to shifts in steroid synthesis during final oocyte maturation (Scott & Sorensen, 1994), and spatially, due to differential release of the pheromone’s components across the gills and in urine (Scott & Ellis, 2007). Both steroid release and EOG studies show that preovulatory females release three key steroid odorants that act via separate, specific, and sensitive (pM threshold) olfactory receptors to induce reproductive responses in males: 17,20b-P, its 20b-sulfated metabolite (17,20b-P-s), and androstenedione (AND) (Scott & Sorensen, 1994; Sorensen et al., 1990; Sorensen, Scott, Stacey, & Bowdin, 1995; Sorensen et al., 2005). As they reach peak synthesis at different times in the preovulatory LH surge, and are released by different routes (17,20b-P and AND across the gills and 17,20b-P-s in urine (Sorensen, Scott, & Kihslinger, 2000; Scott & Ellis, 2007)), these three steroids create a changing mixture in which AND, 17,20b-P, and 17,20b-P-s reach their peak release rates during the early, middle, and late portions of the preovulatory period, respectively (Stacey et al., 1989; Scott & Sorensen, 1994). Based on olfactory detection thresholds from EOG studies, males should detect AND and 17,20b-P only in the immediate vicinity of the female, whereas 17,20b-P-s should be detectable at greater distances (Sorensen et al., 2000; Scott & Ellis, 2007). In the early portion of the female’s LH surge, e.g., the low 17,20b-P : AND ratio of her odor dampens the male LH response (Stacey, 1991) but increases agonistic behavior among males (Poling, Fraser, & Sorensen, 2001). Later, however, increasing 17,20b-P : AND ratios stimulate prolonged following and inspection behaviors among males (Poling et al., 2001) and trigger an LH increase that stimulates milt production and enhances paternity (Dulka et al., 1987; DeFraipont & Sorensen, 1993; Zheng, Strobeck, & Stacey, 1997). In the latter portion of the LH surge, urinary release of 17,20b-P-s induces intense, brief bouts of male chasing and LH increase that likely augments the actions of 17,20b-P on milt production (Sorensen et al., 1995b; Poling et al., 2001). Relatively little is known about how the steroid odorants released by preovulatory females exert their dramatic effects on male behavior and physiology. The increase in male LH following 17,20b-P exposure evidently occurs in response to a reduction of tonic dopaminergic inhibition (Dulka, Sloley, Stacey, & Peter, 1992; Zheng & Stacey,

174

1997). However, nothing is known about how 17,20b-P-s increases male LH, or how AND inhibits the endocrinee gonadal response to 17,20b-P (Stacey, 1991). In competitive spawning, 17,20b-P-exposed males have dramatically greater reproductive success (paternity) than control males (Zheng et al., 1997), an effect that could result from the combined effects of increased competitive behaviors, sperm motility, and sperm release, because 17,20b-P exposure increases all these measures of male reproductive performance (DeFraipont & Sorensen, 1993; Zheng et al., 1997; Hoysak & Stacey, 2008). However, because this reproductive advantage of 17,20b-P-exposed males is also observed in competitive in-vitro fertilizations (Zheng et al., 1997), it seems reasonable to assume that increased sperm quality is a major effect of the preovulatory pheromone. In this regard, it is significant that 17,20b-P, which rapidly increases in male plasma in response to pheromonal 17,20b-P-induced LH increase (Dulka et al., 1987), is known to enhance spermiation, milt hydration, and sperm motility in other fishes (Thomas, 2003; Pankhurst, 2008). Water-borne 17,20b-P not only influences behavior and physiology of male goldfish, but also increases the incidence of ovulation in other female goldfish (Kobayashi et al., 2002); this response is believed to reflect the action of preovulatory female odor on other females because male goldfish appear to release insufficient quantities of 17,20bP and related C21 steroids (Sorensen et al., 2005). This finding suggests 17,20b-P released from females undergoing final oocyte maturation might have triggered the synchronous ovulations observed in laboratory goldfish (Kobayashi et al., 2002), cultured common carp (see Stacey et al., 1994), and crucian carp under natural conditions (Stacey and Olseń, personal observations). The function of such ovulatory synchrony is unknown, but may serve as a predator swamping strategy or reduce the potentially disruptive effects of high male : female sex ratios on female spawning success.

2.1.2. Female postovulatory prostaglandin pheromone When the female goldfish ovulates in the latter half of scotophase, release of the preovulatory steroid pheromone rapidly declines (Stacey et al., 1989; Scott & Sorensen, 1994) and movement of eggs into the oviduct stimulates synthesis of PGF2a, rapidly exerting two critical and simultaneous effects that synchronize the display of male and female spawning behaviors with the presence of ovulated eggs (Stacey & Liley, 1974; Sorensen et al., 1988; 1995a; Kobayashi et al., 2002). Prostaglandin F2a is transported to the brain, where it acts within minutes to stimulate female spawning behaviors (Stacey & Peter, 1979); simultaneously, PGF2a and a metabolite, 15-keto-PGF2a (15K-PGF2a), are released


as a postovulatory pheromone that immediately stimulates male courtship, in turn triggering primer effects that further enhance sperm production (Sorensen et al., 1988; Sorensen, Chamberlain, & Stacey, 1989; Zheng & Stacey, 1996; 1997) (Figure 9.1). Although PGF2a is released both across the gills and in urine, 15K-PGF2a appears to be released almost exclusively in urine pulses, which change in frequency during spawning (Sorensen et al., 1995a; 2000; Appelt & Sorensen, 2007). Prostaglandin F2a and 15K-PGF2a appear to induce similar effects on male behavior and physiology; however, because they are detected by separate olfactory receptor mechanisms, at nM and pM thresholds, respectively (Sorensen et al., 1988), their functions in fact could differ. 15-keto-PGF2a, e.g., might function primarily to alert males to the presence of an ovulated female in the general vicinity, because data from studies of release rate, urinary frequency, and olfactory detection threshold indicate that, during each hour of spawning, a female’s urinations create approximately 40 odor patches, each with an active space of 10 liters (Sorensen et al., 2000; Appelt & Sorensen, 2007). On the other hand, PGF2a may serve to identify a specific ovulated female because it should be detectable only in the immediate vicinity of the female (Sorensen et al., 2000). Although the preovulatory and postovulatory pheromones induce qualitatively similar courtship and LH responses in males, they do so through very different mechanisms (Figure 9.2(a)). Pheromonal 17,20b-P, e.g., acts directly on isolated males to increase plasma LH and milt volume (Sorensen et al., 1989; Fraser & Stacey, 2002). Pheromonal PGFs, on the other hand, evidently induce these effects indirectly, through the sociosexual behaviors they trigger, because PGFs do not increase LH in isolated males (Sorensen et al., 1989) but do increase LH and/or milt when males either are exposed to PGFs in the presence of other males (Sorensen et al., 1989), or spawn with a PGF2a-injected female that is releasing pheromonal PGFs (Zheng & Stacey, 1996). As a result of these differences in pheromonal modes of action, studies of the preovulatory pheromone typically involve simple exposure to waterborne steroids, whereas studies of the postovulatory pheromone involve exposing groups of males to water-borne PGFs, or placing males with sexually active (ovulated or PGF2a-injected) females. A variety of evidence indicates that the goldfish preovulatory and postovulatory pheromones also act through different physiological mechanisms to induce their endocrine and testicular effects in males (Figure 9.2(a)). The preovulatory pheromone appears to stimulate milt production solely through a reduction of tonic dopamine inhibition on pituitary gonadotropes that leads to LH increase and enhanced testicular 17,20b-P production (Dulka et al., 1987; Dulka & Stacey, 1992; Zheng & Stacey,

Chapter | 9

(a)

175


Responses to female cues

17,20β-P

LH

17,20β-P

milt

DA LH

17,20βP

milt

? ?

LH

PGF2 α

milt

PGF2α spawning

(b)

Responses to male cues milt

Stimulated male

LH

? Isolation from males

?

LH

Stimulation

FIGURE 9.2 Schematic models of the multiple mechanisms proposed to regulate milt (sperm and seminal fluid) production in male goldfish. (a) Female pheromones increase milt via mechanisms that either require (4-pregnen-17a,20b-diol-3-one (17,20b-P), preovulatory pheromone) or are associated with (prostaglandin F2a (PGF2a), postovulatory pheromone) male luteinizing hormone (LH) increase. The preovulatory pheromone appears to increase milt only by increasing LH, whereas the postovulatory pheromone also activates an apparently extra-pituitary pathway that rapidly increases milt. DA, dopamine. (b) Exposure to stimulatory males (with high LH) and isolation from inhibitory cues from nonstimulatory males (with low LH) increase milt through longer-latency mechanisms that do not involve LH increase. Reprinted with permission from: Stacey. (2003). Stacey. (2009).

milt

Isolation LH 0

12

milt

24 Hours

1997). For example, pheromonal 17,20b-P-induced LH and milt increase can be blocked either by hypophysectomy or by dopamine D2 receptor agonists, and the latency of the milt effect (approximately three to six hours at 20 C) is very temperature-dependent, as expected of an endocrinemediated process (Dulka et al., 1987; Zheng & Stacey, 1996; 1997). In contrast to the preovulatory pheromone, the postovulatory pheromone appears to act through a mechanism that is distinct and more complex. For example, LH response to the postovulatory pheromone is not blocked by dopamine D2 agonists, and milt responses to the postovulatory pheromone are not blocked by hypophysectomy, and have a shorter latency (< 1 hour at 20 C) that is not temperature-dependent (Zheng & Stacey, 1996; 1997). Further evidence that the two pheromones increase LH through distinct mechanisms is that, even though exposure to supra-threshold concentrations of 17,20b-P induces no further increase in LH (Dulka et al., 1987), combined exposure to 17,20b-P and PGF2a has a synergistic effect on LH secretion (Sorensen et al., 1989). In summary, even our currently rudimentary understanding of the goldfish preovulatory and postovulatory pheromones has revealed an unexpectedly complex suite of physiological and behavioral responses to female odors that evidently have evolved as male tactics for sperm competition. Throughout the periovulatory period, males respond to rapid changes in female odor that both alter male intraand intersexual behavioral interactions, and activate multiple physiological mechanisms that first increase both the quantity and quality of releasable sperm in anticipation of imminent spawning opportunities, and then activate both

endocrine and apparently nonendocrine (i.e., pituitaryindependent) mechanisms that likely function to replenish sperm stores during spawning. Additionally, males regulate releasable sperm not only in response to female odors, but also in response to less-well-characterized cues from males, as briefly discussed in the following section.

2.1.3. The male goldfish pheromone(s) Male goldfish appear to release a hormonal pheromone, likely to be comprised principally of AND, although the function of this pheromone is far less clear than for those produced by females. The key evidence for a male pheromone is that males produce large quantities of AND (Sorensen et al., 2005), that water-borne AND can increase intermale agonistic behavior (Poling & Sorensen, 2001) and inhibit milt production (Stacey, 1991), and that males reduce milt production in the presence of other males that would be releasing large quantities of AND (Stacey, Fraser, Sorensen, & Van Der Kraak, 2001; Fraser & Stacey, 2002). Androstenedione is the most abundant of more than a dozen steroids measured in male goldfish holding water (Sorensen et al., 2005). Androstenedione release by spermiated but sexually inactive males (~50 ngh1) is more than an order of magnitude greater than that released by vitellogenic females (Scott & Sorensen, 1994; Sorensen et al., 2005) and similar to peak levels (~90 ngh1) released by periovulatory females; further, males significantly increase AND release following exposure to pheromonal 17,20b-P (~150 ngh1) or when spawning with PGF2ainjected females (~750 ngh1). Males might be capable of

176

discriminating AND from male and female sources because males release AND in the virtual absence of 17,20b-P and similar C21 steroid odorants (Sorensen et al., 2005), but this has not been investigated. Studies of grouped and isolated male goldfish indicate that males inhibit milt production when exposed to AND from other males. For example, male goldfish evidently inhibit their milt production in the presence of spermiated but sexually inactive males that have basal LH, because their milt volumes dramatically increase after they are isolated (Fraser & Stacey, 2002). This milt increase in the absence of male cues can be blocked by water-borne AND (Stacey, 1991), and is distinct from milt increases in response to female preovulatory and postovulatory pheromones, because it has a longer latency and is not preceded by LH increase (Stacey et al., 2001; Fraser & Stacey, 2002) (Figure 9.2(b)). None-the-less, attempts to use male odor to block milt increase of isolated males have so far been unsuccessful (Fraser & Stacey, 2002) despite experimental designs that should have created detectable AND concentrations (Sorensen et al., 2005), suggesting that, if AND is the factor responsible for the low milt volumes of grouped males, it may be effective only in combination with additional male cues. In addition to reducing milt production in the presence of males with basal LH, male goldfish increase production in the presence of either gonadotropin-injected or 17,20b-Pexposed males (Stacey et al., 2001; Fraser & Stacey, 2002) (Figure 9.2(b)). It is not known what chemical cue(s) males with elevated LH might release to increase milt in other males; however, the results suggest that periovulatory female odor increases milt not only directly, by release of the preovulatory steroid pheromone, but also indirectly, through the males in which they increase LH. If this is in fact the case, then the pressures of sperm competition evidently have selected for mechanisms enabling males to increase both their absolute fertility, by responding to cues from ovulatory females, and their relative fertility, by responding to cues from male competitors. This latter effect is partially comparable to eavesdropping behavior that occurs in visually and acoustically mediated communication networks (McGregor & Peake, 2000; Wisenden & Stacey, 2005). Studies of hormonal pheromones in goldfish and related carps have generated more questions than they have answered. However, it is clear that, despite their outwardly simple (promiscuous and nonparental) mating systems, these fish live in a surprisingly complex social network where hormonal pheromones coordinate multiple functions within and between the genders (Wisenden & Stacey, 2005). Prior to spawning, females release an unidentified recrudescent pheromone that attracts males, which in turn are both sources and receivers of inhibitory cues that suppress milt production. However, when environmental cues trigger a female ovulatory LH surge, release of the


steroidal preovulatory pheromone not only briefly transforms this tonic reciprocal inhibition among males into a positive feedback system, but also triggers ovulatory LH surges in additional females, amplifying the initial stimulus and thereby promoting synchronous final maturation among local conspecifics.

2.1.4. Hormonal pheromones in other cypriniforms Only some of the information derived from hormonal pheromone studies of goldfish and related carps is likely to be applicable to other cypriniforms. It is possible, e.g., that the postovulatory PGF pheromone of goldfish is ubiquitous among cypriniforms, given that all of more than eighty species tested detect PGFs (Stacey & Sorensen, 2006; 2009), and that males spawn with PGF2a-injected females not only in more distantly related species within the cyprinoids (e.g., Java barb (Barbonymus gonionotus) (Cardwell et al., 1995)) but also within the cobitoids (oriental weatherfish loach (M. anguillicaudatus) (Kitamura et al., 1994a)). As in goldfish (Kobayashi et al., 2002), intact but not anosmic male loach display normal courtship and spawning behaviors in the presence of nonovulated females injected with either PGF2a, 15-K-PGF2a, or 13,14dihydro-15-keto-PGF2a (Kitamura et al., 1994a). Nonovulated Misgurnus release negligible amounts of PGFs but, when ovulated, release large amounts of 13,14dihydro-15-keto-PGF2a (>2 mgh1), smaller quantities of PGF2a (~10 ngh1), and virtually no 15-K-PGF2a (Ogata, Kitamura, & Takashima, 1994), consistent with EOG studies (Kitamura et al., 1994a) showing that males are much more sensitive to 13,14-dihydro-15-keto-PGF2a (1 pM detection threshold) than to PGF2a or 15-keto-PGF2a (1 nM detection threshold). Electro-olfactogram screening studies indicate that, unlike PGFs, which likely are detected by all cypriniforms, steroids are detected only by some higher cypriniform taxa, among which the detected compounds can vary greatly (Stacey & Sorensen, 2006; 2009). Indeed, whereas no test steroids were detected by any species from superfamily Cobitoidea (11 species representing all four cobitoid families), steroids appear to be detected by all major lineages within superfamily Cyprinoidea, with the exception of Acheilognathinae (Freidrich & Ko¨rsching, 1998; Pinillos et al., 2002; Lower, Scott, & Moore, 2004; Stacey & Sorensen, 2006; 2009; unpublished). As these EOG screening studies include only a small fraction of extant cypriniforms, and undoubtedly omit steroids that cypriniforms detect, they likely provide a very imperfect picture of cypriniform steroid detection patterns. None-the-less, it seems instructive to ask whether the emerging pattern could offer insights into the evolution of cypriniform hormonal pheromones that might help to focus future studies.

Chapter | 9

177


For example, based on recent molecular studies of cypriniform phylogeny (Wang, Ji, & He, 2007), the most parsimonious explanation for the observed pattern of C21 steroid detection (Figure 9.3) would be that the ability to detect these steroids arose in a common ancestor to the cyprinids, and that the apparent absence of C21 detection in Acheilognathinae is due either to secondary loss or retuning of olfactory C21 receptors. On the other hand, the pattern of C19 detection suggests that the ability to detect these receptors has arisen independently in Leuciscini and Cyprinini (Figure 9.3).

also exhibit positive rheotaxis (facing into a water current) to T (Moore, 1991), but do not increase either plasma hormones or milt volume (Waring, Moore, & Scott, 1996). Moore and Scott (1992) also report that male parr do not normally exhibit an EOG response to 17,20b-P-s, but become extremely sensitive to this steroid (1 pM olfactory threshold) within minutes of a brief (five second) exposure to urine from ovulated females; it will be important to replicate this dramatic finding, as it has not been reported in any other vertebrate. Unfortunately, attempts to determine a biological effect for water-borne 17,20b-P-s on male Atlantic salmon have been unsuccessful (Waring & Moore, 1995; Waring et al., 1996). Despite some important unresolved questions about steroidal odorants in Atlantic salmon, male parr clearly are very sensitive to PGFs (PGF1a and PGF2a) that, as in goldfish (Kobayashi et al., 2002), induce primer effects including increased LH, steroids, and milt (Moore & Waring, 1995; 1996; Olseń, Bjerselius, Mayer, & Kindahl, 2001). Moore and Waring (1996) propose that pheromonal PGFs are released in the urine of ovulated Atlantic salmon because urine and PGFs exert similar priming effects, and ovulated salmon urine contains significant immunoreactive PGF2a. However, Olseń et al. (2001) and Olseń, Johanssen, Bjerselius, Mayer, and Kindahl (2002) propose that ovarian fluid is the major source of the PGF priming pheromone because it has a much higher concentration of PGF2a than does ovulated urine, which, although inducing strong releaser effects, has relatively weak primer effects. In male brown trout, as in Atlantic salmon, ovulated female odor is proposed to contain PGFs that induce endocrine priming effects in males. Further, results of EOG studies (Essington & Sorensen, 1996; Moore & Waring, 1996; Hara & Zhang, 1997; Moore et al., 2002; Laberge & Hara, 2003) agree both on the relative potencies of PGFs (e.g., PGF2a PGF1a > 15K-PGF2a > 13,14dihydro-15K-PGF2a), and that all detected PGFs act through

2.2. Order Salmoniformes Salmoniformes includes only the family Salmonidae, containing the subfamilies Thymallinae (graylings), Coregoninae (whitefishes), and Salmoninae (salmon, trout, and char) (Nelson, 2006); only the latter two subfamilies have been investigated for hormonal pheromones. For coregonins, the only information on hormonal pheromones comes from studies of lake whitefish (C. clupeaformis), which detect two PGF2a metabolites (15K-PGF2a and 13,14-dihydro-PGF2a), which increase locomotory behavior, act through a common receptor mechanism, and induce larger EOG responses in males than in females (Hara & Zhang, 1997; Laberge & Hara, 2003).

2.2.1. Genus Salmo: Atlantic salmon and brown trout Hormonal pheromone studies in Atlantic salmon have focused on the precociously mature male parrda particularly convenient modeldand shown that they detect PGFs and steroids. For example, EOG recordings indicate that precocious males are extremely sensitive to testosterone (T), but only for a brief period prior to spawning, a situation not reported in brown trout or any other fishes. Male parr

C21 Gobionini

C19

+

0

Acheilognathini (Rhodeus ocellatus)

0

0

Tincini

+

0

(Pseudorasbora parva)

1

(Tinca tinca)

Leuciscini

(Rutilus rutilus)

+

+

Xenocypridini

(Ctenopharyngodon idella)

+

0

Cyprinini

(Carassius,Cyprinus)

+

+

Labeoninae

(Epalzeorhynchus frenatus)

+

0

Danioninae

(Danio rerio)

+

0

0

0

Cobitoidea

2

FIGURE 9.3 Relationship between cypriniform phylogeny (Wang et al., 2007) and ability to detect C19 and C21 steroids in electro-olfactogram (EOG) studies. Origin of olfactory sensitivity to C21 steroids may have originated in a common ancestor (*) to Cyprinoidea, whereas olfactory sensitivity to C19 steroids may have originated independently in Leuciscini and Cyprinini. Based on studies by Pinillos et al. (2002)1, Lower, Scott, and Moore (2004)2, and Stacey and Sorensen (2009; unpublished).

178

a single olfactory receptor mechanism. Moreover, as in Atlantic salmon (Olseń et al., 2001; 2002), ovulatory fluid contains far more immunoreactive PGF2a (> 175 ngml1) than does urine (< 5 ngml1), although both increase plasma hormones and milt when added to aquarium water (Olseń et al., 2000; Moore et al., 2002). Despite agreement on relative PGF potencies and receptor number, however, there are inconsistent findings as to whether maturation influences EOG responsiveness. For example, Moore et al. (2002) find that PGFs are detected only by mature brown trout parr (immature parr had no EOG response even at 10 mM), whereas Laberge and Hara (2003) find that undifferentiated juveniles, adult females, and bourgeois males have equivalent EOG responses to PGFs, consistent with the results of Essington and Sorensen (1996) in brown trout and Sveinsson and Hara (2000) in arctic char. Regardless of these inconsistencies, it is clear that water-borne PGFs induce both primer and releaser effects in brown trout. PGF1a and PGF2a increase plasma hormones and milt volume of mature brown trout parr (Moore et al., 2002), as in mature Atlantic salmon parr (Moore & Waring, 1996), and PGFs increase locomotory behavior in bourgeois male trout and induce digging and nest probing (prespawning behaviors) in females (Laberge & Hara, 2003). Hopefully, future studies of Atlantic salmon and brown trout will clarify the origins, release route, and specific pheromonal functions of PGFs, and also confirm reports (Essington & Sorensen, 1996; Hara & Zhang, 1997; Laberge & Hara, 2003) that brown trout do not detect the steroids (T and 17,20b-P-s) proposed to be pheromones in Atlantic salmon; such apparently striking differences between congeners are in marked contrast to the situation in cypriniforms, as discussed above.

2.2.2. Genus Salvelinus (chars) Electro-olfactogram screening studies suggest that chars commonly detect PGFs and are minimally responsive to Etio-g (Essington & Sorensen, 1996; Hara & Zhang, 1997; Laberge & Hara, 2003). However, T-g is reportedly detected by brook char (Salvelinus fontinalis) (Essington & Sorensen, 1996) but not by lake char (S. namaycush) or Arctic char (S. alpinus) (Hara & Zhang, 1997). Electroolfactogram responsiveness to PGF2a is unaffected by gender or maturity in brook and Arctic char (Sveinsson & Hara, 2000), consistent with one study of brown trout (Laberge & Hara, 2003) but in marked contrast to other studies of brown trout (Moore et al., 2002) and Atlantic salmon parr (Moore & Waring, 1995). In Arctic char, PGFs are proposed to function as a male hormonal pheromone that attracts females because males release immunoreactive PGFs and ovulated char are attracted to water-borne PGF2a (Sveinsson & Hara, 2000).


This pheromonal function is the reverse of that suggested for brown trout (Moore et al., 2002) and cypriniforms (Kobayashi et al., 2002), where males are stimulated by postovulatory PGFs, and is particularly surprising given that brook char and brown trout hybridize (Sorensen, Cardwell, Essington, & Weigel, 1995).

2.2.3. Genus Oncorhynchus (Pacific salmon) Electro-olfactogram studies indicate that the olfactory organs of Oncorhynchus spp. are broadly comparable to those of other salmonids in being relatively unresponsive to steroids. For example, the only steroid reported to be detected by chinook salmon (Oncorhynchus tshawytscha), amago salmon (Oncorhynchus rhodurus), and rainbow trout (Oncorhynchus mykiss) is Etio-g (Kitamura, Ogata, & Takashima, 1994b; Hara & Zhang, 1997; Laberge & Hara, 2003; Stacey & Sorensen, 2006; 2009), which is also detected by Atlantic salmon and brown trout, though apparently not by lake whitefish (Essington & Sorensen, 1996; Hara & Zhang, 1997; Laberge & Hara, 2003). Intriguingly, there also are reports that immature rainbow trout detect T (Pottinger & Moore, 1997) and that chinook salmon avoid 17,20b-P (Dittman & Quinn, 1994); it will be important to determine whether these findings can be replicated because they, as with others in Atlantic salmon (Moore & Scott, 1991; 1992), are inconsistent with a general insensitivity to sex steroids among salmonids (discussed by Stacey & Sorensen, 2006; 2009). Unlike the situation with steroid detection, there is a clear distinction between Oncorhynchus and other salmonids with respect to PG detection. For example, EOG studies consistently report not only that coregonins (lake whitefish), Salmo (brown trout, Atlantic salmon), and Salvelinus (brook trout, Arctic char, lake char) detect PGFs (Essington & Sorensen, 1996; Moore & Waring, 1996; Sveinsson & Hara, 2000; Laberge & Hara, 2003), but also that Oncorhynchus (rainbow trout, chinook salmon, and amago salmon) are insensitive to the PGFs with which they have been tested (Kitamura et al., 1994b; Hara & Zhang, 1997; Laberge & Hara, 2003; Stacey & Sorensen, 2006; 2009). Despite their olfactory insensitivity to PGFs, male Oncorhynchus clearly respond to chemical cues from ovulated conspecifics. Ovulatory urine induces both releaser (attraction) and endocrine primer effects in male rainbow trout and masu salmon (Oncorhynchus masou) and, in masu salmon, contains immunoreactive PGF2a (Scott, Liley, & Vermeirssen, 1994; Vermeirssen, Scott, & Liley, 1997; Vermeirssen & Scott, 2001; Yambe, Shindo, & Yamazaki 1999; Yambe & Yamazaki, 2000; 2001a; 2001b; Yambe, Munakata, Kitamura, Aida, & Fusetani, 2003). In masu salmon, the releaser effect of urinary pheromone can be replicated by the tryptophan metabolite, L-kynurenine

Chapter | 9

(Yambe et al., 2006), the first characterized fish sex pheromone that is nonhormonal. It remains to be determined whether L-kynurenine also can induce the primer effects of ovulated masu urine.

2.2.4. Hormonal pheromones and salmonid phylogeny Although potential relationships between steroid detection and salmonid phylogeny are unclear because most EOG recording studies have tested only a small number of steroidal compounds, the emerging pattern of PGF detection raises interesting questions about the origins of salmonid PGF pheromones. From a recently proposed phylogeny (Crespi & Fulton, 2004) (Figure 9.4(a)), PGF detection would appear to be a primitive characteristic of family Salmonidae, because it is seen in the basal whitefishes (subfamily Coregoninae) and all other tested lineages except Oncorhynchus, a pattern that suggests PGFs also are detected by graylings (subfamily Thymallinae). The absence of PGF detection by Oncorhynchus may be due to olfactory receptor loss or retuning, and is reminiscent of the absence of steroid detection by acheilognathin cyprinids (Figure 9.3).

(a)

Oncorhynchus Salvelinus Salmo

?

Thymallus Coregonus

(b)

?

Salmoniformes

Characiformes Gymnotiformes Siluriformes

ostariophysans

?

Cypriniformes

Given that PGF detection may have been present in a common ancestor to salmoniforms, it is interesting to speculate that it also may have been present in a common ancestor to both salmoniforms and osmeriforms, which also detect PGFs (Kitamura et al., 1994b) (Figure 9.4(b)). Similarly, because the common ancestor of ostariophysans likely was sensitive to PGFs, given the widespread detection of PGFs within this taxon, it also is possible that PGF olfactory receptors first originated in a common ancestor to both osteoclupeomorphs and euteleosts (Figure 9.4(b)).

2.3. Order Perciformes Perciform fishes, the dominant vertebrates in both freshwater and marine ecosystems, include more than 10 000 species comprising approximately 40% of living teleosts (Nelson, 2006). Thus, it is surprising that relatively little hormonal pheromone research has been conducted on perciforms, a taxon of immense ecological and economic importance that also provided the first example of a hormonal pheromone in fish (Etio-g in the black goby) (Colombo et al., 1980). Much of our information on perciform hormonal pheromones comes from EOG and other studies of the round goby (N. melanostomus, family Gobiidae) (Murphy et al., 2001; Corkum, Meunier, Moscicki, Zielinski, & Scott, 2008) and cichlids such as Astatotilapia (¼ Haplochromis) burtoni (family Cichlidae) (Cole & Stacey, 2006). Additionally, studies of another perciform, the Eurasian ruffe (Gymnocephalus cernuus, family Percidae) (Sorensen, Murphy, Loomis, Maniak, & Thomas, 2004), clearly illustrate that conclusions drawn from EOG studies alone might be misleading if a species detects only novel hormonal compounds that are not available as test odorants.

2.3.1. The Eurasian ruffe (Gymnocephalus cernuus)

Osmeriformes

?

179


Gonorhynchiformes Clupeiformes FIGURE 9.4 Relationship between salmoniform phylogeny and ability to detect F-series prostaglandins (PGFs) in electro-olfactogram (EOG) studies; salmoniform phylogeny (a) from Crespi and Fulton (2004) and higher order phylogeny (b) from Saitoh, Miya, Inoue, Ishiguro, and Nishida (2003). (a) Sensitivity to PGFs (thick line) may have been present in the common ancestor of salmoniforms, among which thymallins (graylings) are predicted to detect PGFs (thin line); Oncorhynchus spp. appear not to detect PGFs (dotted line). See text for sources of EOG studies.

Eurasian ruffe have recently been introduced to the Laurentian Great Lakes with potentially detrimental effects on native fauna (Gunderson, Klepinger, Bronte, & Marsden, 1998). Studies of potential hormonal pheromones in ruffe (Sorensen et al., 2004) therefore have been undertaken with the expectation that such knowledge could be applied for population control, a strategy that could have general application in fishes, but so far is being pursued only in the sea lamprey, round goby, and common carp (Sorensen & Stacey, 2004; Corkum & Belanger, 2007; Fine & Sorensen, 2008; Sisler & Sorensen, 2008). /If they are exposed to holding water from preovulatory ruffe undergoing oocyte maturation and the plasma surge of 17,20b,21-P (the putative ruffe maturation-inducing steroid (MIS), also known as 20b-S (Pankhurst, 2008)), male ruffe exhibit the same behavioral responses (increased locomotion and social behaviors) that they exhibit to the holding

180

water or urine from nonovulated, 17,20b,21-P-injected females (Sorensen et al., 2004). However, 17,20b,21-P is not responsible for the releaser activity of preovulatory odor, because EOG tests show that ruffe detect neither this compound, nor any of a large number of free and conjugated steroids including those expected to be metabolites of 17,20b-P and 17,20b,21-P (Sorensen et al., 2004). Moreover, preovulatory urine retained its activity after octadecylsilane (C-18) extraction (Sorensen et al., 2004), which removes all known fish pheromones from urine. Sorensen et al. (2004) suggest that, because female ruffe odor has releaser activity only during the preovulatory surge of 17,20b,21-P or following 17,20b,21-P injection, it is likely to be a novel 17,20b,21-P metabolite that is too polar to be extracted with C-18, a conclusion clearly illustrating that EOG screening alone, in the absence of whole-odor bioassays, could greatly underestimate the prevalence of fish hormonal pheromones. The issue is also illustrated by studies of a male urinary pheromone signaling dominance in a cichlid, the Mozambique tilapia (Oreochromis mossambicus), in which EOG testing found no response to a relatively small number of steroids, even though pheromone activity could be demonstrated in a fraction of crude body odor-containing steroid sulfates (Frade, Hubbard, Barata, & Cana´rio, 2002). Further studies (Barata et al., 2008) indicate that the active compound is an uncharacterized sulfated amino-sterol, a chemical category of pheromones previously identified only in the sea lamprey (Sorensen & Hoye, 2007; Fine & Sorensen, 2008).

2.3.2. The black and round gobies The gobies, including as many as 2000 predominantly benthic species, are the largest family of marine fishes. Goby mating systems, even in some pelagic species (Caputo, Mesa, Candi, & Cerioni, 2003), typically involve male territoriality and defense of a nest to which females are attracted for spawning. In pioneering work on the black goby (G. niger), Colombo et al. (1980) reported that the nonspermatogenic but Leydig cell-rich mesorchial gland of the testis synthesizes large amounts of conjugated (glucuronidated and sulfated) 5b-reduced C19 steroids, and that one of these, Etio-g, induces ovulated females to approach and oviposit in empty nests. Since these early studies in black gobies, hormonal pheromone studies have focused on the round goby (N. melanostomus), a recent introduction to the Laurentian Great Lakes that preys on the eggs and young of Great Lakes fishes, and may have contributed to declines of some native species (Corkum, Sapota, & Skora, 2004; Corkum & Belanger, 2007). As with the sea lamprey and Eurasian ruffe (Sorensen & Stacey, 2004), a major incentive for characterizing the hormonal pheromones of the round goby is that the possibility of manipulating or disrupting chemical cues critical for reproductive success


might provide a practical means of biological population control. Initial EOG screening studies (Murphy et al., 2001) showed that round gobies detect not only Etio-g but also more than a dozen conjugated and free C18, C19, and C21 steroids (Figure 9.5); however, unlike cypriniforms, they do not detect known F prostaglandins. Electro-olfactogram cross-adaptation studies indicated that the steroids detected by round gobies are discriminated by four olfactory receptor mechanisms (Figure 9.5(b)), for which the most potent of the tested odorants were estrone (E1), E2-3g, etiocholanolone (Etio), and dehydroepiandrosterone-3sulfate (DHEA-s). Although Murphy et al. (2001) failed to observe any overt reproductive responses following exposure to steroid odors, they fortuitously discovered that exposure to most of the detected steroids increased ventilation (opercular and buccal pumping) frequency, which has been proposed to facilitate odor detection in gobies and other benthic fishes by increasing water flow through the olfactory organ (Nevitt, 1991; Belanger, Corkum, Li, & Zielinski, 2006). Further, even though males and females exhibit equivalent EOG responses to steroid odors, their ventilation responses are sexually dimorphic: i.e., males increase ventilation when exposed to Etio, E1, or E2-3g, whereas females hyperventilate only when exposed to Etio (Murphy et al., 2001) (Figure 9.5(c)). As these hyperventilation responses adapt within 15 minutes, even in continued presence of the odorant, they provide a simple but powerful behavioral cross-adaptation assay that demonstrates two important aspects of the ventilation response to steroid odors. First, male round gobies discriminate the same steroid odors at both the peripheral sensory level (EOG cross-adaptation tests) and central behavioral level (hyperventilation crossadaptation tests) (Figure 9.5(d)) (Murphy et al., 2001). Second, if female gobies receive androgen implants, they not only exhibit the male-typical pattern of hyperventilation in response to Etio, E1, or E2-3g, but also discriminate behaviorally among these steroids in cross-adaptation tests (Murphy & Stacey, 2002) (Figure 9.5(e)). As EOG recordings indicate that olfactory sensory neurons of normal males and females are equally sensitive to steroids (Murphy et al., 2001) (Figure 9.5(a)), these findings therefore suggest that male androgens act on central neural mechanisms to induce the sexually dimorphic hyperventilation response. More recently, the odor of mature male round gobies has been shown to attract vitellogenic females and induce a greater EOG response than the odor of immature males, and the odor of vitellogenic females attracts nonvitellogenic females (Be´langer et al., 2004; Gammon, Li, Scott, Zielinski, & Corkum, 2005). In attempts to characterize the behaviorally active compounds in these goby odors, in-vitro studies have shown that both the testis

Chapter | 9

181


(a)

(c) 8

140

(d) 140

Etiocholanolone

%

mV

Etiocholanolone 10 nM

%

100

1 nM

100

0

Etio

E1

E2-3g

140

Estrone

140 Etiocholanolone 10 nM

%

(b)

Estrone 1 nM

%

Etiocholanolone (Etio) 100

100

Etio E1 E2-3g DHEA-s

0

12

minutes

24

36

0

24

minutes

48

66

(e)

Estrone (E1) Etio E1 E2-3g DHEA-s

140

% Estradiol-3-glucuronide (E2-3g)

Etio E1 E2-3g DHEA-s

100 0 0

50

100

% of initial response

150

minutes

36 Ethanol

Estrone

Estradiol-3glucuronide

72 Etiocholanolone

FIGURE 9.5 Electro-olfactogram (EOG) and behavioral responses (mean standard error of the mean (SEM)) of round gobies to water-borne steroids. (a) EOG responses to etiocholanolone (Etio), estrone (E1), and 17b-estradiol-3-glucuronide (E2-3g) are equivalent in males (filled bars) and females (open bars). (b) EOG cross-adaptation studies indicate that Etio, E1, and E2-3g are each detected by separate olfactory receptor mechanisms. Top: During adaptation to 100 nM Etio, EOG responses to a 10 nM Etio pulse are drastically reduced, whereas responses to pulses of E1, E2-3g, and dehydroepiandrosterone-s (DHEA-s) are unaffected. Middle and bottom: Similar results during adaptation to E1 and E2-3g. (c) Both males (filled circles) and females (empty circles) exhibit a transient increase in ventilation frequency (presented as a percentage of mean pre-exposure frequency) when exposed to 10 nM water-borne Etio (arrow), whereas only males respond to 10 nM E1. (d) Males that are behaviorally adapted by prolonged exposure to 10 nM Etio do not increase their ventilation rate in response to a 10% increase in Etio concentration, but do increase their ventilation rate in response to an equivalent amount of E1. (e) Females implanted with methyltestosterone capsules for 18 days (filled circles) do not increase ventilation when exposed to ethanol vehicle (EtOH) but do increase ventilation when exposed to 1 nM E1, E2-3g, and Etio. In contrast, control females implanted with empty capsules (open circles) respond only to Etio. Redrawn with permission from Murphy et al. (2001) with kind permission from Springer Publishing, from Murphy and Stacey (2002), Stacey and Sorensen (2006), and Stacey and Sorensen (2009) with kind permission from Elsevier, and from Stacey et al. (2003) with kind permission from NRC Press Ottawa.

(Arbuckle et al., 2005) and seminal vesicle (Jasra et al., 2007) convert AND into various 11-oxygenated and 5breduced steroids including Etio and two steroids previously unknown from fishes (11-keto-Etio and 11-keto-Etio-s). However, in recent behavioral assays (Corkum et al., 2008) that used suites of steroids (including 11-keto-Etio and 11-keto-Etio-s) known to be detected by round gobies and/ or produced by mature males, reproductive females were neither attracted nor repelled, whereas nonreproductive females were attracted to a suite of unconjugated steroids and repelled by a suite of conjugated steroids. It seems unfortunate that behavioral studies of goby pheromones have focused predominantly on female

attraction to male odors (Colombo et al., 1980; Be´langer et al., 2004; Gammon et al., 2005; Corkum et al., 2008) given the evidence that goby pheromones mediate additional interactions. Evidence for male response to female pheromones, e.g., comes from classic studies of the frillfin goby (Bathygobious soporator) (Tavolga, 1956) demonstrating that males display courtship behaviors in response to the odor of ovulated females, and from more recent studies (Murphy et al., 2001) showing that male round gobies both detect and respond to steroids such as E1 and E2-3g that presumably are released by females. Further, evidence for intrasexual pheromones in round gobies is provided by reports that females are attracted to female

182

odor (Gammon et al., 2005) and that males both detect and respond to Etio (Murphy et al., 2001) that is produced by males (Arbuckle et al., 2005). An intrasexual pheromone also has been reported in male black gobies, in which agerelated alternate reproductive tactics (Rasotto & Mazzoldi, 2002; Immler, Mazzoldi, & Rasotto, 2004) evidently are associated with the release of distinct odors that may result from differential investment in seminal vesicles, mesorchial glands, and testes. For example, bourgeois males display aggression in response to the odor of bourgeois males, but not in response to the odor of parasitic males (Locatello, Mazzoldi, & Rasotto, 2002), in which the poorly developed mesorchial glands may synthesize insufficient steroid odorants (Rasotto & Mazzoldi, 2002). Being speciose, widely available, and amenable to laboratory manipulations, gobies should become increasingly valuable models for hormonal pheromone research.

2.3.3. Family Cichlidae Cichlids typically exhibit complex social behaviors mediated by multiple sensory modalities. Although cichlid visual communication has received the most attention (e.g., Oliveira, Lopes, Carneiro, & Canario, 2001; Seehausen et al., 2008), it is clear that interactions with conspecifics also are mediated through audition (Amorim, Simo˜es, Fonseca, & Turner, 2008) and olfaction (Frade et al., 2002; Miranda, Almeida, Hubbard, Barata, & Cana´rio, 2005; Barata, Hubbard, Almeida, Miranda, & Cana´rio, 2007; Barata et al., 2008). No cichlid pheromone has been chemically identified, although Barata et al. (2008) propose that the urine of male Mozambique tilapia contains an amino-sterol pheromone that signals male dominance status to females. In addition, EOG studies in the African mouthbrooder A. burtoni (Cole & Stacey, 2006) provide clear evidence for pheromonal steroids. As in the round goby (Murphy et al., 2001), A. burtoni does not detect a variety of F- and E-series prostaglandins, but does detect diverse C18, C19, and C21 steroids (Cole & Stacey, 2006). However, the round goby detects both conjugated and unconjugated steroids, whereas A. burtoni appears to detect only conjugated (glucuronidated and sulfated) steroids. Further, the results of EOG crossadaptation and binary mixture tests indicate that A. burtoni’s olfactory organ contains five olfactory receptor mechanisms capable of discriminating the nature and position of the conjugate: e.g., 3-glucuronide; 17-glucuronide; 3-sulfate; 17-sulfate; 3,17-disulfate. Although there is no evidence that A. burtoni releases any of the steroids it is able to detect, or that the steroid odorants induce a biological response, the presence of a complex pheromone system in this species is indicated both by the number and variety of conjugated steroids that are discriminated (Cole & Stacey, 2006), and by the fact that


these conjugates are very likely to be released in controlled urine pulses (Appelt & Sorensen, 2007; Barata et al., 2007; Scott & Ellis, 2007). Pinheiro, Souza, and Barcellos (2003) report that male Nile tilapia (Oreochromis niloticus) increase milt volume and sperm motility following exposure to water-borne 17,20b-P, and also preferentially court females that have been injected with 17,20b-P, a behavioral response that is blocked by anesthetic-induced anosmia (Souza, Lucion, & Wassermann, 1998). These findings are surprising, however, since EOG studies indicate not only that cichlids appear to detect only conjugated steroids but also that other Oreochromis species (O. mossambicus, O. tanganicae, and O. aureus) do not detect 17,20b-P (Frade et al., 2002; Cole & Stacey, 2006; Stacey, 2009; unpublished). Cichlids appear to be an ideal taxon in which to explore the evolution of hormonal pheromone function because they are widely distributed and speciose, and there is increasing consensus on the phylogenetic relationships of higher order cichlid taxa (Koblmu¨ller, Sefc, & Sturmbauer, 2008). Family Cichlidae consists of four major clades (Sparks & Smith, 2004; Genner et al., 2007), the basal Etroplinae and Ptychochrominae found in India and Madagascar, and two derived sister groups, the African Pseudocrenilabrinae and the New World Cichlinae. A. burtoni is found within the ‘modern haplochromines’ (Salzburger, Mack, Verheyen, & Meyer, 2005), the remarkably diverse (> 1500 spp.) crown group of pseudocrenilabrines that displays a propensity for explosive speciation not seen in other cichlid lineages. It therefore seems reasonable to ask not only whether the ability of A. burtoni to detect conjugated steroids (Cole & Stacey, 2006) is shared by, and perhaps unique to, other haplochromines, but also where in the evolutionary history of cichlids these olfactory sensitivities arose, and whether they might have facilitated the explosive speciation of haplochromines by promoting reproductive isolation through enhancement of chemically mediated discrimination of conspecifics. To begin to answer these questions, I am conducting EOG screening studies to determine the extent and pattern of steroid detection among cichlids, and to identify taxa that could serve as model species. Although preliminary, the results (Stacey, 2009; unpublished) clearly suggest that the cichlids’ olfactory responsiveness to conjugated steroids arose in African pseudocrenilabrines after their divergence from the New World cichlines. In EOG recordings of over seventy species from all four major cichlid clades, no steroids were detected by etroplines, ptychochromines, or cichlines, whereas at least some conjugates were detected by all tested pseudocrenilabrines (> 50 spp.) with the exception of Heterochromis multidens, the basal pseudocrenilabrine species (Stacey, 2009; unpublished) (Figure 9.6). Moreover, as in A. burtoni, other pseudocrenilabrines detect only

Chapter | 9

Haplochromis Aulonocara Lethrinops Cyphotilapia

3-sulph

Lamprologus Chalinochromis Spathodus

Cichlidae

183


Pseudocrenilabrinae Cichlinae Ptychochrominae Etroplinae

17-gluc 17-sulph 3,17-disulph

Tilapia Steatocranus Hemichromis Heterochromis Crenicichla Geophagus

No steroids detected

Paratilapia Katria Paretroplus

Labridae

3-gluc

FIGURE 9.6 Cichlid phylogeny (based on Sparks & Smith, 2004; Salzburger, Mack, Verheyen, & Meyer, 2005) and the origins of olfactory sensitivity to steroid conjugates. Astatotilapia burtoni detects a range of conjugated steroids (Cole & Stacey, 2006) that appear to be detected by five olfactory receptor mechanisms responding to 3-glucuronides (3-g), 3-sulfates (3-s), 17-glucuronides (17-g), 17-sulfates (17-s), and 3,17-disulfates (3,17-di-s). Ongoing electro-olfactogram (EOG) studies (Stacey, 2009; unpublished) suggest detection of steroid conjugates arose in the pseudocrenilabrines and proceeded by progressive accumulation of olfactory receptor mechanisms. Representative sample species listed alphabetically are: Aulonocara stuartgranti, Chalinochromis euchilus, Crenicichla sp., Cyphotilapia frontosa, Geophagus brasiliensis, Astatotilapia burtoni, Hemichromis stellifer, Heterochromis multidens, Katria katria, Lamprologus congoensis, Lethrinops auritus, Paratilapia polleni, Paretroplus maculatus, Spathodus erythrodon, Steatocranus cassuarius, Tilapia discolor.

(not tested)

conjugated steroids, which in virtually all cases are the same as those detected by A. burtoni (Cole & Stacey, 2006; unpublished). However, it is very important to note that, unlike the situation with A. burtoni (Cole & Stacey, 2006), investigations of other pseudocrenilabrines have not yet included EOG cross-adaptation studies to determine which of the detected steroid conjugates acts through separate olfactory receptor mechanisms, and thus might be expected to be perceived as distinct odors. None-the-less, on the assumption that the steroid receptor mechanisms proposed for A. burtoni are not only homologous but also still functionally similar to those of less derived lineages, the most parsimonious interpretation of the current EOG findings is that putative steroidal pheromone function in pseudocrenilabrines has evolved primarily by a sequential accumulation of the five olfactory receptor mechanisms proposed for A. burtoni (Cole & Stacey, 2006). For example, the 3-glucuronide receptor mechanism of A. burtoni appears to be the primitive cichlid glucuronide receptor and to have arisen in an ancestor common to basal West African hemichromines (Hemichromus spp.) and tilapiines (Oreochromis, Pelvicachromis, Steatocranus, and Tilapia spp.). Not only do these taxa appear to possess only a single olfactory receptor mechanism sensitive to 5b,3aglucuronides such as Etio-g, but also an apparently similar 3-glucuronide mechanism is seen in all more-derived pseudocrenilabrines (Figure 9.6). Similarly, sensitivity of A. burtoni to 17-glucuronides, 17-sulfates, and 3,17-disulfates appears to have arisen in a common ancestor of eretmodines (Spathodus erythrodon) and more derived pseudocrenilabrines (Figure 9.6), a group corresponding to the MVhL lineage of Takahashi, Terai,

Nishida, and Okada (2001) which includes seven tribes that have been subject to EOG screening (Eretmodini, Lamprologini, Ectodini, Cyphotilapiini, Limnochromini, Haplochromini, Tropheini) and two that have not (Perissodini, Cyprichromini). All tested MVhL species detect 17b-glucuronides (E2-17g), 17b-sulfates (T-s), and 3,17b-disulfates (E2-3,17s) (Cole & Stacey, 2006; Stacey, 2009; unpublished). The origin of the fifth A. burtoni receptor mechanism (detecting 3-sulfates) is less clear (Figure 9.6). Although 3-sulfates are detected by all tested cyphotilapiines (Cyphotilapia), haplochromines (Pseudocrenilabrus, Pundamilia, Lethrinops, etc.), and tropheins (Tropheus, Lobochilotes), some tribes have not been examined (perissodines, cyprichromines), and, within ectodines, some species (Xenotilapia ornatipinnus) are responsive whereas others (Ophthalmotilapia nasuta) are not. Currently, it seems that species responsive to 3-sulfates correspond to the C-lineage proposed by Clabaut, Saltzburger, and Meyer (2005; also see Koblmu¨ller et al., 2008). In addition to demonstrating an apparently simple pattern of steroid conjugate receptor increase during pseudocrenilabrine evolution, EOG screening studies have revealed what appear to be novel conjugate receptors restricted to several taxa. For example, Tylochromis sudanensis, the only tylochromine yet tested, is similar to species in other basal West African tribes (hemichromines, tilapiines) in detecting 3-glucuronated steroids such as Etio-g, but also detects two types of conjugates not known to be detected by other Africans: T-g and 17,20b-P-g. As T-g is a 17b-glucuronated androstene, the Tylochromis receptor detecting this steroid is likely distinct from the

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17b-glucuronide receptors of the MVhL cichlids, which respond only to 17b-glucuronated estratrienes (E2-17g). Detection of 17,20b-P-g by Tylochromis is particularly notable because it is the only known instance of 20bglucuronide detection by any cichlid; however, a similar olfactory novelty appears to have arisen in X. ornatipinnus, the only cichlid known to detect a 21-sulfated steroid (e.g., 4-pregnen-11b,17,21-diol-3,20-dione-21-s ¼ cortisol-21s). Clearly, it will be important to determine whether such apparently novel conjugate receptors are restricted to T. sudanensis and X. ornatipinnis or are common to these and related genera. Although these EOG studies have barely begun to sample from the rich diversity of African cichlids, they already are contributing to an understanding of how hormonal pheromone systems might arise and evolve. First, it is clear that the apparently complex steroidal pheromone system of A. burtoni did not arise de novo in the relatively recent haplochromine lineage, but rather had its origins much earlier, near the base of pseudocrenilabrine diversification (see Genner et al. (2007) and Koblmu¨ller et al. (2008) for discussion of aging of cichlid diversification). Second, the pattern of olfactory receptor accumulation during pseudocrenilabrine evolution has identified taxa that should be informative for clarifying the order in which steroid conjugate receptors have arisen. For example, it seems likely that the apparently simultaneous appearance of three receptors (17-glucuronide, 17-sulfate, and 3,17-disulfate) in the MVhL lineage is an artifact of insufficient sampling, and that future work on more basal tribes (Bathybatini, Trematocarini, Boulengerochromini) might reveal sequential accumulation. Finally, by tracing both the origins and phylogenetic distributions of olfactory sensitivities to steroid conjugates, these cichlid EOG studies clearly demonstrate that hormonal pheromones can be both homologous and analogous: to cite just one example, not only is there widespread detection of 3-glucuronides (Etio-g) throughout pseudocrenilabrines, but the ability to detect these conjugates also has arisen independently in other perciforms (Murphy et al., 2001) and in salmoniforms (Laberge & Hara, 2003).

3. HORMONAL PHEROMONES AND THE ISSUE OF SPECIES SPECIFICITY Given the key physiological and behavioral responses that sex pheromones induce, and the likelihood that it would be maladaptive to exhibit these responses when exposed to heterospecific odors, it is understandable that sex pheromones are commonly expected to be species-specific. Further, from this perspective, it also might be expected that sex hormones would be poor candidates for


pheromonal function, for the simple reason that the highly conserved biosynthetic pathways for steroids and prostaglandins should place severe constraints on the chemical diversity that enables specificity. It therefore is no small conundrum that hormonal pheromones not only appear to be widespread in many speciose taxa (e.g., cypriniforms, characiforms, salmonids, siluriforms, cichlids (Stacey & Sorensen, 2002; 2009)), but also might be extremely similar, if not identical, among at least some closely related species (Irvine & Sorensen, 1993; Narayanan & Stacey, 2003). As the species specificity of hormonal pheromones is virtually unexplored, the following brief discussion is offered in the hope of stimulating future research, the success of which relies heavily on selection of appropriate model species. Reduction in fitness resulting from response to heterospecific odors is likely the major selective pressure for evolution of a recognizable conspecific odor; selection of study species, therefore, should be based on some understanding of the likelihood and fitness costs of heterospecific responses, and the mechanisms by which the resulting selection pressure could alter specificity. In order for maladaptive responses to heterospecific odor to select for specificity, a species must be in functional reproductive sympatry, such that individuals enter the active space of heterospecifics when physiologically capable of pheromone-induced responses. A key factor determining the frequency of functional reproductive sympatry, therefore, is the size of the active spaces created by the released hormonal odorants of conspecifics; these typically should be small, given that hormones and hormonal precursors and metabolites are in low concentrations even prior to release. Although swimming speed and water flow will influence the shape and size of pheromonal active spaces, odorous filaments will result from continuous and passive release of unconjugated steroids (AND and 17,20b-P) across the gills, whereas odorous patches will be produced by pulsatile release of prostaglandins (15K-PGF2a) and some conjugated steroids (17,20b-P-s) in urine (Scott & Ellis, 2007). In goldfish, e.g., the size of active spaces can be roughly estimated because rates of hormone release and olfactory detection thresholds are known (Scott & Sorensen, 1994; Sorensen et al., 2000; 2005). In spawning males, e.g., AND release is estimated to create relatively large active spaces (Table 9.2) (Sorensen et al., 2005), presumably due to high plasma AND levels, although these have not been measured. Release of AND and 17,20b-P by periovulatory females, however, generates relatively small active spaces detectable only in their immediate vicinity (Table 9.2), although their pulsatile release of 17,20b-P-s and 15KPGF2a should create urinary patches detectable at somewhat greater distances. Overall, the apparently small active spaces of most goldfish hormonal pheromones likely result from living in bisexual aggregations that obviate the

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TABLE 9.2 Theoretical active spaces of pheromonal compounds released across the gills and in the urine of periovulatory female goldfish (F) and mature male goldfish that are either sexually inactive (M) or spawning (Msp)1 Sex

Compound

Release route

Release duration (h)2

Peak release (ng.h1)

EOG threshold (log M) 11

Active space (l.h1)3

Reference

M

AND

G

continuous

~50

~10

~17

5

Msp

AND

G

variable4

~750

~1011

~250

5

F

AND

G

6

~90

~1011

~30

6

F

17,20b-P

G

12

~60

~5 1012

~35

6

F

17,20b,21-P

G

3

~60

~5 1011

~3

6

F

17,20b-P-s

U

9

~65

~1011

~8 3

6e8

F

17,20b,21-P-s

U

9

~150

~5 1011

~8 1

6e8

F

15K-PGF2a

U

variable4

~750

~5 1012

~40 11

8e10

1

After Stacey (2009), with permission. The time that the compound should be detectable, when neither the source fish nor the receiver are moving, and the odor is being dispersed by diffusion only. Defined as the calculated volume of water in which a pheromone of known release rate will be detectable, and calculated as the quantity of pheromone (moles) released per h divided by the EOG detection threshold of the receiver. Calculations based on data from Scott and Sorensen (1994) and Sorensen et al. (1988, 2000, 2005). 4 Determined by the time (typically several hours) required to complete oviposition. 5 Sorensen et al. (2005). 6 Scott and Sorensen (1994). 7 Sorensen et al. (1995b). 8 Appelt and Sorensen (2007). 9 Sorensen et al. (1988). 10 Sorensen (unpublished results). 2 3

necessity of detecting conspecifics at a distance, and considerably larger active spaces might therefore be anticipated in species such as gobies, in which pheromones from nesting males are proposed to attract females (Colombo et al., 1980; Gammon et al., 2005). None-theless, evidence that active spaces of hormonal pheromones in natural waters are quickly reduced by degradation (Sorensen et al., 2000) and by interactions with humic acids (Mesquita, Canario, & Melo, 2003) suggests the frequency of functional reproductive sympatry might often be insufficient to select for specificity, although this very important question has not been addressed. Even when functional reproductive sympatry occurs, however, evolution of specificity would be expected only if fitness costs were sufficient. In a semelparous female salmonid, e.g., ovulating in response to heterospecific odor could result in total reproductive failure and thereby exert strong selective pressure for specificity. However, in an iteroparous male cyprinid such as the goldfish, nonspecificity might well persist if increases in milt production in response to heterospecific odor occurred infrequently, and thus had negligible impact on fitness. Although it is relatively straightforward to envisage scenarios wherein selection for species specificity should occur, it should not be assumed that there will necessarily

be a mechanism enabling selection to lead to specificity. For example, we have proposed (Wisenden & Stacey, 2005; Stacey & Sorensen, 2006; 2009) three evolutionarily related conditions (‘ancestral,’ ‘spying,’ and ‘communication’) to conceptualize how released hormonal products influence conspecific interactions (Figure 9.7). All fish are, or have been, in the ancestral (prepheromonal) condition in which the hormones and related compounds released by originators are not detected by conspecifics. This ancestral condition progresses to spying if conspecifics evolve the ability to respond adaptively to (i.e., benefit from) a released hormonal product (now termed a pheromonal cue), and then might progress to communication if there is a mechanism that enables the receiver’s response to exert selective pressure for specialization in production and/or release of the cue (now termed a signal). Thus, the key distinction between communication and spying is that, in communication, both pheromone producer and receiver are specialized for pheromonal functions, whereas in spying such specializations are restricted to receivers (Figure 9.7). As discussed more fully by Sorensen and Stacey (1999) and Wisenden and Stacey (2005), the major reason this hypothetical scenario is relevant to studies of pheromonal specificity is that, because fitness benefits (or loss thereof) accrue to both signaler and receiver in communication, but

186


COMMUNICATION (originator becomes signaler) RESPONSE Receiver

Signaler

Receiver’s response selects for signal specialization

SIGNAL Change in chemical and/or its release

SPYING (change in receiver) Receiver

Originator CUE

Evolution of detection and response. Receiver benefits.

Chemical and its release unchanged.

ANCESTRAL Receiver

Originator CHEMICAL

No mechanisms for detection or response.

Chemical released to the water.

FIGURE 9.7 Proposed stages in the evolution of pheromonal communication: ancestral stage, where hormones and metabolites are released but not detected by conspecifics; spying stage, where receivers evolve specializations for detecting hormonal compounds but originators are not specialized for their release; communication stage, in which both originator (now a signaler) and receiver are specialized for transfer of chemical information. Redrawn from Stacey and Sorensen (2002; 2006) with kind permission from Elsevier; from Wisenden and Stacey (2005) with kind permission from Cambridge University Press; and from Stacey and Sorensen (2008) with kind permission from Science Publishers.

can be restricted to only the receiver in spying, selection pressures for specificity might be expected to be different in the two conditions. For example, where a spying pheromone benefits only the receiver, specificity might be achieved only by changes in what receivers detect, because there may be no mechanism for the receiver’s response to select for specificity by altering pheromone release. In communication, a potential result of receivers’ selective pressure on signalers is signal amplification (the goby mesorchial gland might be an example (Arbuckle et al., 2005)), which could increase the number of species in functional reproductive sympatry by increasing pheromonal active space; such alteration of pheromone release would not occur in spying, in which, by definition, only receivers are specialized for pheromonal function. From the preceding discussion, the likelihood that hormonal pheromones typically have small active spaces implies that many fishes may experience no fitness cost from the use of hormonal pheromones that are not speciesspecific. Additionally, the fact that fishes not only release and detect a large number of steroid metabolites (Sorensen & Scott, 1994; Arbuckle et al., 2005; Sorensen et al., 2005; Scott & Ellis, 2007) but also have extraordinary olfactory abilities to discern small differences in steroid structure (Sorensen et al., 1988; 1990; 1995a; 1995b; Murphy et al.,

2001; Cole & Stacey, 2006) suggests that evolving speciesspecific steroidal pheromones might in fact not be problematic. The potential for species-specific prostaglandin pheromones is much less clear, however, because so little is known about prostaglandin metabolism in fishes (Sorensen et al., 1995a). Electro-olfactogram studies have convincingly demonstrated that fishes detect a surprising diversity of steroidal compounds, including unconjugated and conjugated forms not only of estrogens (17b-estradiol (E2)) (Murphy et al., 2001), androgens (11-ketotestosterone (11-KT)) (Cardwell et al., 1995), and MISs (17,20b-P) (Sorensen et al., 1990), but also of their precursors and metabolites (Scott & Sorensen 1994; Murphy et al., 2001; Narayanan & Stacey 2003; Cole & Stacey, 2006) (Figure 9.8). C21 steroids in particular might be especially important for generating species-specific odors because, with the exception of C17 glucuronidation, they can be conjugated with sulfate or glucuronide at all five carbons (C3, C11, C17, C20, and C21) that commonly bear oxygen groups. When this variability is combined with the various configurations (5b,3a-; 5b,3b-; 5a,3a-; and 5a,3b-) resulting from A-ring reduction, the diversity of C21 metabolites that fishes could potentially release is enormous. Despite revealing great differences in the hormones and related compounds that fishes detect, EOG studies have provided virtually no evidence that such differences result from selection for conspecific recognition (or heterospecific avoidance). Indeed, it is particularly striking that, whereas distantly related taxa such as mochokid catfishes (Narayanan & Stacey, 2003), cichlids (Cole & Stacey, 2006; unpublished results), goldfish (Kobayashi et al., 2002), and gobies (Murphy et al., 2001) detect markedly different steroids, remarkably similar hormonal compounds can be detected by closely related species (congeners and related genera), among which the potential for heterospecific interaction would be greatest (Irvine & Sorensen, 1993; Stacey & Sorensen, 2006; 2009). In characiform tetras (Astyanax, Hemigrammus, Hyphessobrycon), e.g., all of more than a dozen species tested detected only E2-3s (Cardwell & Stacey, 1995). Similarly, common carp, crucian carp, and goldfish detect and respond to very similar steroid and prostaglandin odorants (Irvine & Sorensen, 1993; Stacey et al., 1994; Bjerselius et al., 1995; Olseń et al., 2006), possibly accounting for their ability to hybridize in natural conditions (Ha¨nfling, Bolton, Harley, & Carvalho, 2005). Even where congeners detect different steroids, this need not imply selection for specificity. For example, it seems likely that in the speciose (> 100 species) African catfish genus Synodontis (family Mochokidae), patterns of steroid sensitivities (similar among sympatrics but different among allopatrics) result from geographic dispersal and subsequent divergence rather than from selection for

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187


(a) 17b -Estradiol metabolites E2-3-glucuronide

E2-17-glucuronide

E2-3,17-disulphate

O Gl

O

OH

Gl O Neogobius melanostomus Oreochromis aureus

O SO3

O3S O

Haplochromis burtoni Tropheus moori

Haplochromis polystigma Cheilochromis euchilus

(b) Androgen-related metabolites Etiocholanolone

Androsterone-3-sulphate

O

HO

O3S O

H

Testosterone-17-sulphate O

O

H Tyrranochromis nigriventris Lethrinops auritus

Neogobius melanostomus Synodontis ocellifer

FIGURE 9.8 Diversity of steroid hormone metabolites that induce electro-olfactogram (EOG) responses in fishes. Representative sample species are megalopids (Megalops cyprinoides), cyprinids (Carassius auratus, Cyprinus carpio, Epalzeorhynchos frenatus), siluriforms (Synodontis ocellifer and macrops), gobies (Neogobius melanostomus), and cichlids (Cheilochromis euchilus, Cynotilapia afra, Astatotilapia burtoni, Hemichromis elongatus, Lethrinops auritus, Nimbochromis polystigma, Oreochromis aureus, Pelvicachromis pulcher, Pundamilia nyererei, Tropheus moori, Tylochromis sudanensis, Tyrranochromis nigroventris, Xenotilapia ornatipinnis).

SO3

O Pundamilia nyererei Cynotilapia afra

(c) MIS-related metabolites 4-pregnen compounds 17,20b P-20-glucuronide

17,20b P-20-sulphate CH3 SO3 HC O OH

CH3 Gl HC O OH

O Tylochromis sudanensis Carassius auratus

4-Pregnen-11b,21-diol3,20-dione-21-sulphate

O Epalzeorynchos frenatus Cyprinus carpio

CH2 SO3 HC O OH

O Xenotilapia ornatipinnis

pregnan compounds 5b-Pregnan3a,17,20b-triol

5b-Pregnan-3a,17-diol20-one-3-glucuronide CH3 O CH OH

CH3 HO CH OH

HO

H

Megalops cyprinoides Synodontis macrops

5a-Pregnan-3b,20b diol-3-sulphate

Gl O

H

Hemichromis elongatus Pelvicachromis pulcher

O

O3S O

CH3 CH

H

Xenotilapia ornatipinnis

specificity (Narayanan & Stacey, 2003). All of more than a dozen tested Synodontis species detect a suite of unconjugated C21 (5b,3a-pregnan) and C19 (5b,3a-androstan) steroids that varies geographically: i.e., although all species detect the same 5b,3a-androstan steroids, species from the north and east (Lake Nyassa) detect a different suite of 5b,3a-pregnans than do species from the central African Zaire River region. These findings in tetras, carps, and catfish suggest that, once a hormonal pheromone function evolves, it is likely to be retained by descendent species with little if any modification in the compounds detected. This is perhaps most forcefully illustrated by the African

pseudocrenilabrine cichlids, in which the ability to detect 5b,3a-glucuronides (5b-pregnan-3a,17-diol-20-one-3aglucuronide; 5b-androstan-3a-ol-17-one-3a-glucuronide ¼ Etio-g) evidently originated in basal west African groups such as the hemichromines and tylochromines (Stacey, 2009; unpublished) (Figure 9.4) and has been retained in descendent pseudocrenilabrines for at least 30 million years (Genner et al., 2007). Electro-olfactogram studies showing that related species can detect the same or very similar hormonal compounds (Irvine & Sorensen, 1993; Narayanan & Stacey, 2003) seem inconsistent with the finding that, in response to

188


whole body odor, many fishes clearly discriminate conspecifics from heterospecifics (Stacey, Kyle, & Liley, 1986; Sorensen & Stacey, 1999; Sisler & Sorensen, 2008) and also display surprising patterns of conspecific and heterospecific preference (Wong, Fisher, & Rosenthal, 2005). Possible resolutions to this apparent discrepancy might simply be that: (1) fishes are able to discriminate subtle differences in ratios of a common hormonal mixture; (2) the hormonal products conferring specificity have not yet been included in EOG screening studies and (3) the pheromones contain both nonspecific hormonal components and species-specific nonhormonal components. The research required to resolve these issues will not be trivial, but the bioassay-guided high performance liquid chromatography (HPLC) fractionation approach first used to identify the novel bile acid pheromone of sea lamprey (Fine & Sorensen, 2008) and more recently applied to cichlid pheromones (Barata et al., 2008) should serve as both a guide and incentive for exploring such fundamental questions of hormonal pheromone function.

ACKNOWLEDGEMENTS The author acknowledges a career’s worth of generous funding from the Natural Sciences and Engineering Research Council of Canada (NSERC) and is very grateful to Alison Murray for helpful comments on portions of an earlier draft.

ABBREVIATIONS 11-KT 15K-PGF2a 17,20b-P 17,20b-P-s AND C18 DHEA-s E1 E2 E2-17g E2-3,17s E2-3g EOG Etio Etio-g GTH HPLC LH MIS PGF PGF1a PGF2a T T-g T-s

11-ketotestosterone 15-keto-PGF2a 4-pregnen-17a,20b-diol-3-one 20b-sulfated metabolite of 17,20b-P Androstenedione Octadecylsilane Dehydroepiandrosterone-3-sulfate Estrone 17b-estradiol 17b-estradiol-17b-glucuronide 17b-estradiol-3,17b-disulfate 17b-estradiol-3-glucuronide Electro-olfactogram Etiocholanolone Etiocholanolone-3b-glucuronide Gonadotropin High performance liquid chromatography Luteinizing hormone Maturation-inducing steroid F2-series prostaglandin Prostaglandin F1a Prostaglandin F2a Testosterone Testosterone glucuronide Testosterone sulfates

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Van Den Hurk, R., & Resink, J. W. (1992). Male reproductive system as sex pheromone producer in teleost fish. J. Exp. Zool., 261, 204e213. Vermeirssen, E. L. M., & Scott, A. P. (2001). Male priming pheromone is present in bile, as well as urine, of female rainbow trout. J. Fish Biol., 58, 1039e1045. Vermeirssen, E. L. M., Scott, A. P., & Liley, N. R. (1997). Female rainbow trout urine contains a pheromone which causes a rapid rise in plasma 17,20b-dihydroxy-4-pregnen-3-one levels and milt amounts in males. J. Fish Biol., 50, 107e119. Wang, X., Li, J., & He, S. (2007). Molecular evidence for the monophyly of East Asian groups of Cyprinidae (Teleostei: Cypriniformes) derived from the nuclear recombination activating gene 2 sequences. Mol. Phylogenet. Evol., 42, 157e170. Waring, C., & Moore, A. (1995). F-series prostaglandins have a pheromonal priming effect on mature male Atlantic salmon (Salmo salar) parr. In F. W. Goetz, & P. Thomas (Eds.), Proceedings of the Fifth International Symposium on the Reproductive Physiology of Fish (pp. 255e257). Austin, TX: Fish Symposium 95. Waring, C. P., Moore, A., & Scott, A. P. (1996). Milt and endocrine responses of mature male Atlantic salmon (Salmo salar L.) parr to water-borne testosterone, 17a,20b-dihydroxy-4-pregnen-3-one 20-sulfate, and the urines from adult female and male salmon. Gen. Comp. Endocrinol., 103, 142e149. Wisenden, B. D., & Stacey, N. E. (2005). Fish semiochemicals and the evolution of communication networks. In P. McGregor (Ed.), Animal Communication Networks (pp. 540e567). London, UK: Cambridge University Press. Wong, B. B. M., Fisher, H. S., & Rosenthal, G. G. (2005). Species recognition by male swordtails via chemical cues. Behav. Ecol., 16, 818e822.


Yamazaki, F. (1990). The role of urine in sex discrimination in the goldfish Carassius auratus. Bull. Fac. Fisher. Hokkaido Univ., 41, 155e161. Yambe, H., & Yamazaki, F. (2000). Urine of ovulated female masu salmon attracts immature male parr treated with methyltestosterone. J. Fish Biol., 57, 1058e1064. Yambe, H., & Yamazaki, F. (2001a). A releaser pheromone that attracts methyltestosterone-treated immature fish in the urine of ovulated female trout. Fish. Sci., 67, 214e220. Yambe, H., & Yamazaki, F. (2001b). Species-specific releaser effect of urine from ovulated female masu salmon and rainbow trout. J. Fish Biol., 59, 1455e1464. Yambe, H., Munakata, A., Kitamura, S., Aida, K., & Fusetani, N. (2003). Methyltestosterone induces male sensitivity to both primer and releaser pheromones in the urine of ovulated female masu salmon. Fish Physiol. Biochem., 28, 279e280. Yambe, H., Kitamura, S., Kamio, M., Yamada, M., Matsunaga, S., Fusetani, N., et al. (2006). L-Kynurenine, an amino acid identified as a sex pheromone in the urine of ovulated female masu salmon. Proc. Natl. Acad. Sci., 103, 15370e15374. Yambe, H., Shindo, M., & Yamazaki, F. (1999). A releaser pheromone that attracts males in the urine of mature female masu salmon. J. Fish Biol., 55, 158e171. Zheng, W., & Stacey, N. E. (1996). Two mechanisms for increasing milt volume in male goldfish. J. Exp. Zool., 276, 287e295. Zheng, W., & Stacey, N. E. (1997). A steroidal pheromone and spawning stimuli act via different neuroendocrine mechanisms to increase gonadotropin and milt volume in male goldfish (Carassius auratus). Gen. Comp. Endocrinol., 105, 228e235. Zheng, W., Strobeck, C., & Stacey, N. E. (1997). The steroid pheromone 4-pregnen-17a,20b-diol-3-one increases fertility and paternity in goldfish. J. Exp. Biol., 200, 2833e2840.

Chapter 10

Reproduction in Agnathan Fishes: Lampreys and Hagfishes Stacia A. Sower* and Hiroshi Kawauchiy *

University of New Hampshire, Durham, NH, USA, y Laboratory of Molecular Endocrinology, Sendai, Miyagi, Japan

SUMMARY This chapter summarizes reproduction and the latest findings on reproductive endocrinology in two representatives of the most ancient lineage of vertebrates: agnathans. Modern vertebrates are classified into two major groups: the gnathostomes (jawed vertebrates) and the agnathans ( jawless vertebrates). The agnathans are classified into two groups: myxinoids (hagfish) and petromyzonids (lampreys), while the gnathostomes constitute all the other living vertebrates including the bony and cartilaginous fishes and the tetrapods. During the past two decades there have been rapid advances in our knowledge of the structure and function of reproductive hormones in lampreys. However, key elements of the endocrine reproductive system in hagfish have yet to be elucidated. Here we present a summary of our current knowledge of reproductive endocrinology in these basal vertebrates.

1. INTRODUCTION The relatedness of the hagfish and lampreys, the two most basal extant vertebrates, can be judged from certain key anatomical features that they share, but they differ profoundly in other ways. Both groups lack jaws, paired fins, a third semicircular canal, and neural arches over the spinal cord. Both have a single median gonad and a distinctively thin, flat adenohypophysis that is broken up into lobules. They differ in the basic structure of the intestine, in the germ layer origins of the olfactory organ and adenohypophysis, and in the numbers of eggs that they produce. In terms of their relatedness to higher vertebrate taxa, the lampreys share features, or resemble, these higher taxa much more than do the modern hagfishes. The hypothalamicepituitary system is considered to be a vertebrate innovation and seminal event that emerged prior to or during the differentiation of the ancestral agnathans (Sower, Freamat, & Kavanaugh, 2009). Reproduction in vertebrates is controlled by a hierarchically organized endocrine system. In spite of the very diverse patterns of


lifecycles, reproductive strategies, and behaviors, this endocrine system is remarkably conserved throughout the gnathostome lineages. To date, biochemical, molecular, immunocytochemical, and functional studies on the structure and function of the gonadotropin-releasing hormones (GnRHs) and glycoprotein hormone (GpH) in lampreys have established that, similar to all other vertebrates, the lampreys have a hypothalamicepituitaryegonadal (HPG) axis and that there is a high conservation of the mechanisms of GnRH action (Sower et al., 2009) (Figure 10.1). On the other hand, in hagfish there seemed to be a lack of, or poor regulation of reproduction by, hypothalamicepituitary peptides (Sower & Gorbman, 1999). Until recently, inadequate experimental support had confounded the presence and possible function of sex-steroid hormones and hypothalamic neurohormones such as GnRH (Powell, Kavanaugh, & Sower, 2005). Hagfish studies from the past few years now show seasonal changes in GnRH, gonadal steroids, estradiol (E2), and progesterone (P4), corresponding to gonadal reproductive stages along with the putative identity of a functional corpus luteum (Powell, Kavanaugh, & Sower, 2004; Kavanaugh, Powell, & Sower, 2005; Powell, Kavanaugh, & Sower, 2005; 2006). Recent studies also have shown immunoreactive (ir)-gonadotropin (GTH) in the anterior pituitary of the hagfish and demonstrated that the GTHtype cell predominated in the pituitary in adults with developing gonads (Miki, Shimotani, Uchida, Hirano, & Nozaki, 2006; Nozaki, Shimotani, & Uchida, 2007). The cDNA of one GTH has been reported in brown hagfish (Paramyxine atami) at an international meeting (Honda, Uchida, Shimotani, Moriyama, & Nozaki, 2006). Hagfishes mostly inhabit a deep marine environment that is relatively free of circadian, or even seasonal, changes. Lampreys, on the other hand, either are totally freshwater dwellers or they at least reproduce in freshwater after periods in the sea, migrating between the two. How much the modern hagfishes and lampreys represent or 193

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FIGURE 10.1 (a) Schematic representation of the hypothalamicepituitary axes in adult sea lampreys (Petromyzon marinus). Distribution of immunoreactive (ir)-gonadotropin-releasing hormone (GnRH) containing cell bodies (circles) and fibers (broken lines) in approximate midsagittal planes in the brain of the sea lamprey. Distribution of ir-gonadotropin (GTH)b in the adenohypophysis of lampreys. Data regarding GnRH in lampreys are from Nozaki, Tsukahara, and Kobayashi (1984); Nozaki and Gorbman (1985); King, Sower, and Anthony (1988); Kavanaugh, Nozaki, and Sower (2008). Data regarding GTHb in lampreys are from Sower et al., 2006. (b) Schematic representation of the hypothalamicepituitary axes in adult Atlantic hagfish (Myxine glutinosa). Distribution of ir-GnRH containing cell bodies (circles) and fibers (broken lines) in approximate midsagittal planes in the brain of hagfish. Distribution of ir-GTH (X) in the adenohypophysis of hagfish, respectively. Data regarding GnRH in hagfish are from Braun, Wicht, and Northcutt (1995); Sower, Nozaki, Knox, and Gorbman (1995). Data regarding GTH in hagfish from Miki, Shimotani, Uchida, Hirano, and Nozaki (2006); Nozaki, Shimotani, and Uchida (2007). ADW, anterior dorsal wall of the neurohypophysis; AH, adenohypophysis; CT, connective tissue; DHN, dorsal hypothalamic nucleus; DT, dorsal thalamus; DW, dorsal wall of the neurohypophysis; Hyp, hypothalamus; IPN, interpeduncular nucleus; IR, infundibular recess; NH, neurohypophysis; PI, pars intermedia; PPD, proximal pars distalis; PON, preoptic nucleus; PPN, posterior part of the preoptic nucleus; POC, postoptic commissure; PoON, postophic nucleus; OC, optic chiasma; RPD, rostral pars distalis; T, optic tectum; VW, ventral wall of the neurohypophysis.

reflect the ancient forms from which they evolved 500 million years ago is a question that still concerns evolutionary biologists and that is far from being answered.

2. HAGFISH REPRODUCTION Reproductive patterns in the two agnathan groups, as far as they are known, are clearly divergent. Lamprey reproduction is a visible and thoroughly described event. It follows ritual nest building and mating behavioral phenomena that can be witnessed in freshwater streams and lakes. Hagfish reproduction and early development, because of their inaccessibility, have so far escaped observation and can only be inferred. Lampreys die after fecundation of their eggs. Key elements of the reproductive system have not been elucidated in hagfish. However, there is now evidence showing that some species of hagfish may have seasonal reproductive cycles. The Japanese hagfish (Eptatretus burgeri) is the only known species of hagfish that has a regular annual reproductive cycle; it undergoes an annual migration from shallow waters during late October to July to deeper waters in August and September during the spawning season (Kobayashi, Ichikawa, Suzuki, & Sekomoto, 1972; Ichikawa, Kobayashi, & Nozaki, 2000; Nozaki, Ominato, Gorbman, & Sower, 2000). The seasonal migration and seasonal development of gonads were examined in E. burgeri near the Misaki Marine Biological Station of the University of Tokyo during the period from October 1970

to October 1975 (Ichikawa et al., 2000; Nozaki et al., 2000). In these studies, the reasons for the seasonal migration of E. burgeri did not appear to be simply related to reproduction but to many other factors such as food supply and water temperature (Ichikawa et al., 2000). There are reports on the seasonal movements of two other species of hagfish (Eptatretus deani and Eptetratus stoutii); however, the reasons for these migrations are not known (Martini, 1998). In terms of the gonads, there was no difference in the annual growth curves of developing eggs or testicular development in Japanese hagfish sampled in shallow vs. deep water; however, the developing eggs were smallest in October and largest in September (Nozaki et al., 2000). There is now evidence from recent reproductive studies that the Atlantic hagfish Myxini glutinosa also may have a seasonal reproductive cycle (Powell et al., 2004; Kavanaugh et al., 2005; Powell et al., 2005). Steroid concentrations from in-vitro incubations of gonads have been shown to be correlated with brain concentrations of GnRH with development and maturation of gonadal tissues in Atlantic hagfish captured monthly from the Atlantic Ocean (Powell et al., 2004; Kavanaugh et al., 2005). These data showed an annual cycle of brain GnRH as well as E2 and progesterone P4 production from the gonads in medium and large hagfish. The increase in in-vitro E2 production was also correlated with an increase in the number of maturing eggs in female hagfish. ‘Brown bodies’ were identified in the ovaries and were shown to have both biochemical and

Chapter | 10

Reproduction in Agnathan Fishes: Lampreys and Hagfishes

morphological properties consistent with a functional corpus luteum (Powell et al., 2006). These data support a seasonal reproductive cycle in the Atlantic hagfish.

2.1. The Hagfish Gonad The gonads of both hagfish sexes are single median structures suspended from the ventral side of the gut, which in turn is suspended from the dorsal body wall (Brodal & Fange, 1963; Sower & Gorbman, 1999). The gonadal elements are contained in a membrane that extends the full length of the body cavity. The testis occupies the extreme posterior end of the membrane, and it is relatively small. The ovary comprises most of the length of the membrane. Hagfishes were once considered functional hermaphrodites but the frequency of hermaphrodites in most hagfish populations is not known (Powell et al., 2005). Hermaphrodites were rarely observed in a study on the Japanese hagfish (Ichikawa et al., 2000). In general, little information is available on sex ratios and sex determination in hagfishes. Based on recent studies, about 40 to 50% of medium-sized Atlantic hagfish sampled had gonads with distinct male testicular tissue in the posterior region and ovarian tissue in the anterior region (Powell et al., 2004). From part of this same study, 58% of all hagfish studied (n ¼ 1080) from all size classes contained only female gonad tissue, 41% were hermaphrodites, and 0.05% were males with no ovarian tissue present. These authors proposed that a certain percentage of Atlantic hagfish may indeed be functional hermaphrodites since a small percentage of the identified hermaphroditic adult hagfish were found with large oval eggs and mature sperm (Powell et al., 2004). Even though this is a high percentage of hermaphroditism, it is not yet known whether these Atlantic hagfish are functional hermaphroditesdi.e., releasing mature eggs and sperm simultaneously for self-fertilization. The question of whether Atlantic hagfish are hermaphroditic and the possible reproductive significance of this group remain unknown. It is hypothesized that Atlantic hagfish have the capability of being functional hermaphrodites as well as functioning separately as males and females. These authors speculated that this may be a strategy that has helped the Atlantic hagfish and perhaps other hagfish species to survive over millions of years in sometimes extreme oceanic conditions such as a nutrient-limited environment (Powell et al., 2005).

2.1.1. Sex differentiation All post-hatching, smaller hagfishes are female. In the most thoroughly studied species (E. stouti) the gonad contains only small nonvitellogenic oocytes until the animal attains a length of about 20 cm. Thus, sex differentiation is considered to be progynous (Gorbman, 1990).

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In differentiation of male animals larger than 20 cm, the oocytes degenerate and the posterior end of the gonadal membrane develops spermatic follicles. In females, the posterior part of the gonad involutes and the long anterior section develops vitellogenic oocytes and, finally, mature eggs. It is of interest that many populations of hagfishes have been reported to consist of variable ratios of males to females, in which the females are more numerous. Thus, it is possible that environmental factors influence sex differentiation.

2.1.2. The ovary The germinative tissue of the adult hagfish ovary is restricted to the edge of the membranous gonad (Brodal & Fange, 1963; Fernholm, 1975; Dodd & Dodd, 1985). Numerous small oocytes move dorsally in the membrane as they grow. It is uncertain whether these growing, rounded oocytes actually move or whether their apparent movement is due to growth in depth of the membrane. As the oocytes grow they gradually assume an oval shape, and finally a spindle shape. In E. stouti, when the oocytes reach a length of about 4.5 mm, many become atretic, and only about 25 to 30 of them continue growing as active vitellogenesis begins (Brodal & Fange, 1963). It would appear that, even after selection of 25 to 30 eggs for the completion of maturation has begun, production of smaller oocytes continues; however, all of these are destined for atresia. When the selected eggs have reached a length of more than 25 mm, the follicular epithelium (granulosa) of each egg thickens and begins to secrete a shell, which, of course, precludes further growth of the egg (Fernholm, 1975). The follicular epithelium at either end of the egg becomes thrown into a complexly folded pattern and secretes the two crowns of anchor-shaped hooks. A hole in the shell remains (the micropyle) to allow sperm entrance for fertilization. The remarkably complex morphogenetic properties of the hagfish granulosa epithelium deserve more study. Corpora lutea in nonmammalian vertebrates secrete mainly P4, thought to be involved in the retention of eggs and downregulation of vitellogenin synthesis (Chieffi & Baccari, 1998). The most ancient vertebrate that was known to have a functional corpus luteum is the dogfish Squalus acanthias (Tsang & Callard, 1987). Brown bodies, hypothesized to be corpora lutea, had been observed by scientists for over 100 years in the gonad of an even more ancient lineage of vertebrate, the hagfishes (Walvig, 1963; Patzner, 1978). However, data in support of brown bodies acting as corpora lutea had consisted mainly of observational studies. Powell et al., 2006 proposed that hagfish have functional corpus lutea-like structures that produce P4, based on electron and light-microscopy studies that showed P4 production from the ‘brown bodies,’ as opposed to

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oocytes. These authors further hypothesized that one possible ancestral function of P4 in hagfishes is an increase in vascularization of the ovary, and that this function was established at an early stage of evolution in vertebrates (Powell et al., 2006).

2.1.3. The testis The gonadal membrane in the posterior few abdominal body segments, in which the hagfish testis forms, is much shorter than it is in the ovarian area (Brodal & Fange, 1963). One lateral surface of the membrane in the testicular area becomes thickened and folded and this epithelial area is the source of the spermatic follicles that comprise the testis. Since the testis is so small compared to the ovary, earlier authors pointed out that the volume of sperm produced must be too small to permit a system of external fertilization in the open seawater. Fertilization must be in an enclosed space to achieve a high enough concentration of spermatozoa to assure success. This is especially so because of limited access to the egg through the micropyle and the unusually small number (less than 30) of mature eggs provided by females. However, no observations of the fertilization process have been reported, and so this suggestion awaits verification.


regulation of adenohypophysis secretion by neurohormones diffusing from the neurohypophysis. The adenohypophysis of hagfishes appears to be minimally, if at all, functional (Sower & Gorbman, 1999); its cells contain very few stainable granules. Attempts to show whether the usual pituitary tropic hormones can be extracted or immunochemically stained have given equivocal results. Recently, three different types of adenohypophysial cells were revealed in the pituitary of the brown hagfish (P. atami) by means of immunohistochemistry in combination with lectin histochemistry (Miki et al., 2006). The major findings of these studies suggested that GTH and corticotropin (ACTH) are major pituitary hormones in hagfishes. The cell type that was ir to ovine luteinizing hormone (LH) predominated in the adenohypophysis in adults with developing gonads and thus appeared to be involved in the regulation of gonadal functions. This is further substantiated by a more recent but unpublished observation in which identification of GTH has been reported in the brown hagfish (P. atami) (Honda et al., 2006). Sower and Gorbman (1999) hypothesized that the extent to which the deep-sea habitat lacks photic or temperature/seasonal/cyclic clues suggests that there is less of a need for a functional HPG axis in hagfishes. The extent of pituitary involvement in hagfish reproduction will require further study.

2.2. Secondary Sexual Characteristics Hagfishes have no gonadoducts and they present no externally differentiated sexual features (Brodal & Fange, 1963). Tsuneki and Gorbman (1977) pointed out that in male E. burgeri a mucus gland near the cloacal pore is larger in males than in females. It has been claimed also that the thread cells in the slime glands of the skin could be considered secondary sex features because the threads that they form (in both sexes) may be used to enclose and protect the developing eggs on the sea floor.

2.3. The Hypothalamicepituitaryegonadal (HPG) Axis of Hagfishes 2.3.1. Neurohypophysis and adenohypophysis The hagfish neurohypophysis is a flattened sac-like structure that contains nerve endings of neurons that originate in nuclei of the hypothalamus. These neurons contain two forms of ir GnRH (Braun, Wicht, & Northcutt, 1995; Sower, Nozaki, Knox, Gorbman, 1995). The adenohypophysis is a thin layer of follicular groups of cells that is separated by connective tissue from the neurohypophysis (Figure 10.1). This arrangement of the hypothalamicepituitary structures resembles that of lampreys, even in lacking any direct nervous or vascular connection between the two. It is an anatomical arrangement structure that would seem to favor

2.3.2. Reproductive hormones Gonadotropin-releasing hormone is the major hypothalamic neurohormone involved in mediating reproductive activity in vertebrates (Kavanaugh, Nozaki, & Sower, 2008). Gonadotropin-releasing hormone is a peptide composed of ten amino acids that is released from the hypothalamus by appropriate internal and external cues and then travels to the anterior pituitary, where it stimulates the release of GTHs. The GTHs, LH, and follicle-stimulating hormone (FSH) enter the bloodstream and act on the gonads to stimulate steroidogenesis and induce the maturation of eggs or sperm. Until recently, evidence supporting the presence of GTH in hagfishes was not conclusive. Matty, Tsuneki, Dickhoff, and Gorbman (1976) identified only limited abnormalities in the testes and ovaries of 150 hypophysectomized hagfish (E. stoutii) during a seven-month study. Gametogenesis appeared to be unaffected by hypophysectomy, suggesting that the hagfish gonad was independent of hypophysial gonadotropic control. However, Patzner and Ichikawa (1977) observed a decrease in the number of follicles containing spermatocytes and only a few follicles containing spermatids in hypophysectomized E. burgeri when compared with sham-operated hagfish. These results suggest that the development of the gonad in this hagfish is under hypophysial gonadotropic control.

Chapter | 10


However, in hagfishes there seems to be a lack of, or poor regulation of, reproduction by hypothalamice pituitary peptides (Sower & Gorbman, 1999). Inadequate experimental support has not conclusively shown the presence and possible function of sex steroid hormones and hypothalamic neurohormones such as GnRH. In fact, GnRH has not been isolated from any hagfish. In two studies using antisera to lamprey GnRH-III from Dr. Sower’s laboratory, as well as other GnRH antisera from Dr. Northcutt’s laboratory, ir GnRH was detected in the brain of the Atlantic hagfish and Pacific hagfish (E. stouti) (Braun et al., 1995; Sower et al., 1995). These data suggested the presence of a lamprey GnRH-III-like molecule in the hagfish brain and that hagfishes may have reproductive control mechanisms that are similar to other vertebrates. Based on this evidence that GnRH is present in hagfish, ir-GnRH was measured in Atlantic hagfish sampled seasonally (Kavanaugh et al., 2005). Seasonal changes of GnRH correlated with a seasonal cycle of steroid hormones and gonadal development proposed by Powell et al. (2004) and Kavanaugh et al. (2005). These data suggest that an active neuroendocrine axis is responsible for the reproductive cycle of the hagfish. However, full determination of the HPG axis in hagfish will require the identification of the native GnRHs and other reproductive hormones to fully assess the extent of the HPG axis.

3. LAMPREY REPRODUCTION 3.1. The Lamprey Gonad 3.1.1. Lifecycle There are approximately 40 species of lamprey, which are classified as parasitic or nonparasitic. Lampreys are semelparous; i.e., they spawn only once in their lifetime, after which they die. Sexual maturation is thus a synchronized process coordinated with the life stages of the lamprey. The reproductive cycle of the sea lamprey Petromyzon marinus is the best known and will serve as a model for lamprey reproduction. The sea lampreys begin their lives in fresh water as blind, filter-feeding larvae. After approximately five to seven years in freshwater streams, metamorphosis occurs and the ammocoetes become free-swimming, sexually immature lampreys, and migrate to the ocean. Lamprey metamorphosis is a highly synchronized and programmed process that involves major physiological and morphological change (reviewed in Youson & Sower, 2001). In the parasitic sea lamprey, sexual maturation is a seasonal, synchronized process. During the long parasitic ocean phase, gametogenesis progresses slowly. In males, spermatogonia proliferate and develop into primary and secondary spermatocytes; in females, vitellogenesis occurs.

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After approximately 15 months at sea, lampreys return to freshwater streams and undergo the final maturational processes, resulting in production of mature eggs and sperm, spawning, and then death. In nonparasitic species, gonadal differentiation and metamorphosis occur together. Shortly after the end of the maturation of gametes in parasitic lamprey species, spawning occurs and the lampreys die.

3.1.2. Development of larval gonads The gonad in both sexes of sea lampreys is unpaired, medial, and suspended from the dorsal wall of the body cavity by means of a mesentery containing connective tissue. Lampreys are among the vertebrates, including teleost fish, that have no intraperitoneal genital ducts. Hardisty (1971; pp 295e360) has summarized the most important stages in the development of the larval gonads as follows: “(1) The appearance during gastrulation of the primordial germ cells and their migration to the gonadal site below the aorta. (2) The development of a median genital ridge extending caudally from the pronephric region. At this stage, the gonadal rudiment contains a relatively small number of germ cells and is covered by a peritoneal epithelium. (3) After hatching, for periods varying from six months to over two years, the undifferentiated gonad shows comparatively little further development and throughout this stage the germ cells divide only slowly, if at all, remaining solitary or arranged in groups of two to four secondary deuterogonia.1 (4) This is succeeded by a period of more rapid mitotic division, during which the germ cells tend to remain together to form cell nests or cysts. (5) The onset of the meiotic prophase occurs both in isolated germ cells and in the cysts. These meiotic cells may proceed to cytoplasmic growth or may undergo atresia in the earlier stages of prophase. (6) Oogenesis occurs to a variable extent in the great majority, if not all, ammocoete gonads, but in the future ovaries it tends to be more synchronous, leading eventually to a gonad containing no germ cells other than oocytes in the cytoplasmic growth phase. (7) In the differentiation of the male gonads, a large proportion of the germ cells undergo degeneration in the earlier stages of the meiotic prophase, oocytes which survive to the cytoplasmic growth phase are eventually eliminated by atresia. Because of this extensive degeneration of germ cells, the testis is reduced in size and contains only small numbers of 1

‘Deuterogonia’ refers to groups of germ cells that will differentiate or undergo atresia.

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residual germ cells. It is from these undifferentiated elements that nests of primary spermatogonia are developed by renewed mitotic divisions either shortly before, or during metamorphosis.” Following these stages either during or after metamorphosis, gametogenesis continues.

3.1.3. The ovary During the final maturational stages in the female sea lamprey, there is an intense, rapid gonadal growth period a few weeks before spawning (Hardisty, 1971). As female sea lampreys enter fresh water, the oocytes are ready for spawning with most of the cytoplasm filled with yolk platelets. The germinal vesicle of the egg is located peripherally, the vitelline membrane has doubled in thickness, and the theca has begun to separate from the vitelline membrane. At ovulation, the thecal follicle cells separate from the oocytes. Some time before ovulation actually occurs, the granulosa cells become distended with secretory materials (gelatinous material) and show a maximal degree of separation of one from the other. This gelatinous material, containing granules of acid mucopolysaccharides, is not liberated until ovulation, when the granulosa cells disintegrate and leave remnants attached to the basal pole of the oocyte, forming the adhesive layer that causes the eggs to stick to the substrate when they are finally shed. The number of eggs produced by each lamprey varies with species. The smallest number of eggs is produced by Mordacia praecox, which matures about 450 eggs at the time of spawning (Hardisty, 1971). The largest known numbers of eggs is produced by the largest lamprey species, P. marinus, estimated to be from 124 000 to 260 000 (Hardisty, 1971).

3.1.4. The testis In male parasitic lampreys returning to the rivers in early autumn or late spring, only primary spermatogonia are present in the testes (Hardisty, 1971). The testes grow by mitotic division of spermatogonia. At the end of this growth phase, spermatogonia are transformed into spermatocytes. This stage appears to last for several months in river lampreys and for a few weeks in sea lampreys. The final stages of spermatogenesis occur during the last few weeks before spawning and the testicular ampullae become filled with mature spermatozoa. Prior to spawning, the testicular ampullae break down and the sperm are liberated into the body cavity before exiting via genital pores.

3.2. Secondary Sexual Characteristics In lampreys, both sexes develop secondary sexual characteristics during the final weeks of reproduction and spawning

activity (Hardisty, 1971). In both sexes, the fins enlarge at the time of prespawning sexual maturity and the two dorsal fins become continuous. In addition, there are swollen cloacal labia and curvature of the tail, and an erectile ‘penis’ develops in the cloacal region of the male. The tail of the female curves sharply upwards at sexual maturity and in the male there is a slight downward curvature.

3.3. The Hypothalamicepituitaryegonadal (HPG) Axis of Lampreys 3.3.1. Neurohypophysis and adenohypophysis In lampreys, the neurohypophysis is not as highly developed as in hagfishes (Gorbman, 1983). It consists of a thin anterior section that is the floor of the diencephalon. The posterior part of the neurohypophysis is somewhat thickened and is the terminating neurohemal structure for neurons whose cell bodies are in the preoptic part of the hypothalamus and thus similar to a pars nervosa (Gorbman, 1983). The adenohypophysis (anterior pituitary) is much better differentiated than in hagfishes and is differentiated into the pars intermedia and pars distalis. Neither the hagfishes nor lampreys have the anatomical equivalent of a median eminence. The median eminence in gnathostomes is the neurohemal structure that conveys the neurohormones to the adenohypophysis via portal blood vessels. Of all vertebrates, only the agnathans and the teleosts lack this portal vascular system for transferring regulatory peptides from the brain to the adenohypophysis. Based on earlier experiments (Nozaki, Fernholm, & Kobayashi, 1975; Tsukahara, Gorbman, & Kobayashi, 1986; Nozaki, Gorbman, & Sower, 1994) in which substances of varying molecular size were injected into the third ventricle of hagfish and lampreys and shown to diffuse rapidly from the third ventricle, through the neurohypophysis, to the pars distalis, it was proposed by Nozaki et al. (1994) that the extant agnathans had a ‘diffusional median eminence’ (p 390). In other words, neurosecretory peptides such as GnRH are able to diffuse from the brain to the pituitary to regulate its secretory activity.

3.3.2. Gonadotropin-releasing hormone (GnRH) The GnRH family currently includes 28 isoforms, 15 and 13 from representative vertebrate and invertebrate species, respectively (Kavanaugh et al., 2008; Tsai & Zhang, 2008; Zhang, Tello, Zhang, & Tsai, 2008). To date, two to three isoforms have been identified in representative species of all classes of vertebrate (Gorbman & Sower, 2003; Guilgur, Moncaut, Canario, & Somoza, 2006; Kah et al., 2007; Kavanaugh et al., 2008; Okubo & Nagahama, 2008). Early analyses by Grober, Myers, Marchaterre, Bass, and Myers (1995) and Fernald and White (1999) suggested that there

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are three paralogous groups of GnRH in gnathostome brains. In recent reviews of GnRH and respective receptors, various scenarios on the phylogenetic relationships among just the gnathostomes’ GnRHs and receptors have been suggested (Guilgur et al., 2006; Kah et al., 2007; Okubo & Nagahama, 2008). We have shown from an extensive analysis of all available cDNA GnRH sequences, including the eight cloned lamprey GnRH-III cDNAs, that the vertebrate GnRHs are grouped into four paralogous lineages: GnRH 1, 2, 3, and 4 (Silver, Kawauchi, Nozaki, & Sower, 2004). We based our analysis on phylogenetic analysis, function, neural distribution, and developmental origin (Silver et al. 2004). This phylogenetic analysis confirmed the earlier model of three GnRH lineages and, in addition, showed that lamprey GnRH-I and -III form a fourth group. The identification of two more invertebrate mollusk GnRHs and one in Annilid has suggested a fifth grouping of GnRHs (Aplyia, octopus, limpid, annilid GnRH) in invertebrates and supports the fourth group, consisting of lamprey GnRH-I and -III (Tsai & Zhang, 2008; Zhang et al., 2008). The newly discovered lamprey GnRH-II offers a new paradigm of the origin of the vertebrate GnRH family (Kavanaugh et al., 2008). Likely due to a genome/gene duplication event, an ancestral gene gave rise to two lineages of GnRHsdthe gnathostome GnRH and lamprey GnRH-II. The gene duplication events that generated the different fish and tetrapod paralogous groups likely took place within the gnathostome lineage, after its divergence from the ancestral agnathans. Lamprey GnRH-I and -III (type 4) can be identified now as paralogous homologs of gnathostome GnRH and lamprey GnRH-II group (type 2), resulting from a duplication within the lamprey lineage. This implies that there probably was a genome/gene duplication event that gave rise to all forms of vertebrate GnRH affecting the common ancestor of lamprey and gnathostome isoforms. Lampreys are the most basal vertebrates for which there are demonstrated functional roles for multiple GnRH neurohormones that are involved in pituitaryereproductive activity. Both lamprey GnRH-I and -III have been shown to induce steroidogenesis and spermiation and/or ovulation in adult sea lampreys (Sower, 2003). In lampreys undergoing metamorphosis, there is an increase in brain lamprey GnRH-I and -III that coincides with the acceleration of gonadal maturation (Youson & Sower, 1991). In immunocytochemical studies, both lamprey GnRH-I and -III immunoreactions can be found in the cell bodies in the rostral hypothalamus and preoptic area in larval and adult sea lampreys (King, Sower, & Anthony, 1988; Wright, McBurney, Youson, & Sower, 1994; Tobet, Nozaki, Youson, & Sower, 1995; Nozaki et al., 2000). It has been suggested that, in the larval stage, most of the ir-GnRH is lamprey GnRH-III (Tobet et al., 1995). Recently, a cDNA encoding a novel GnRH was cloned from the sea lamprey (Kavanaugh et al., 2008). As with all

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other vertebrate GnRHs, the lamprey GnRH-II is highly conserved, with 10 amino acids and conserved C- and Ntermini. Similar to GnRH-II expression in gnathostomes, lamprey GnRH-II has been shown to be widely distributed and expressed in a number of tissues. In the brain, lamprey GnRH-II has been shown to be located in the preoptic area/ hypothalamus by in-situ hybridization and immunocytochemistry. In contrast to the type 2 GnRH (chicken GnRHII) generally found in the midbrain of gnathostomes and generally acting as a nonhypothalamic hormone in gnathostomes, lamprey GnRH-II may have a role as a third hypothalamic form in lampreys. Further studies will be needed to assess the actions of GnRH-I, -II, and -III on the synthesis and/or release of lamprey GTH to understand the differential activities of these three GnRHs.

3.3.3. Gonadotropin-releasing hormone receptor (GnRH-R) At the vertebrate adenohypophysis, GnRH action is mediated through high-affinity binding with the GnRH receptor (GnRHR), a class A or rhodopsin-like seven-transmembrane Gprotein-coupled receptor (GPCR). Gonadotropin-releasing hormone receptors have been classified typically as type-1 (without a carboxy-terminal (C-terminal) tail; GnRH-R 1) and type-2 (with a C-terminal tail; GnRH-R 2) (Silver, Nucci, Root, Reed, & Sower, 2005; Guilgur et al., 2006; Kah et al., 2007). The GnRH-R 1 is unique among all GPCRs in that this receptor lacks the highly conserved intracellular C-terminal tail. A full-length transcript encoding a functional GnRH-R 1 has been isolated and cloned from the pituitary of the sea lamprey (Silver et al., 2005). The cloned receptor retains the conserved structural features and amino acid motifs of other known GnRH-Rs and notably includes a C-terminal intracellular tail of ~120 amino acids. The lamprey GnRH receptor was initially shown to activate the inositol triphosphate (IP3) signaling system; stimulation with either lamprey GnRH-I or lamprey GnRH-III led to dose-dependent responses in COS7 (mammalian cell line) cells transiently transfected with lamprey GnRH-R. Expression of the receptor transcript was demonstrated in the pituitary and testes using reverse transcription polymerase chain reaction (RT-PCR), while in-situ hybridization showed expression and localization of the transcript in the proximal pars distalis of the pituitary (Silver et al., 2005). The phylogenetic placement and structural and functional features of this GnRH receptor suggested that it is representative of an ancestral GnRH receptor.

3.3.4. Gonadotropin (GTH) In gnathostomes, the GpH family consists of two pituitary GTHs (LH and FSH), one chorionic GTH (CG), and one thyrotropin (TSH). Each of these GpHs consists of

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a heterodimer composed of an a- and a b-subunit. The asubunit is common within a single species (Kawauchi et al., 1989). The b-subunits are homologous and convey hormone specificity (Kawauchi et al., 1989; Swanson, 1991; Huhtaniemi, 2005). Two GTHs have been identified in all taxonomic groups of gnathostomes (Suzuki, Kawauchi, & Nagahama, 1988; Kawauchi et al., 1989; Querat, Sellouk, & Salmon, 2000; Querat et al., 2004; Huhtaniemi, 2005). To date, only one GTH has been identified in jawless vertebrates (agnathans) (Sower et al., 2006). We have identified the first and perhaps only GTH b-like protein by cDNA cloning in sea lampreys (Sower et al., 2006). As the sea lamprey GTH b protein is a clear out-group compared to those of the LH and FSH family based on phylogenetic analysis, we had proposed that an ancestral GpH gave rise to only one GTH in lampreys and to the GpH family that, in turn, gave rise to LH, FSH, and TSH during the early evolution of gnathostomes. A fifth heterodimeric GpH (after FSH, LH, TSH, and CG) was discovered in 2002 and termed ‘thyrostimulin’ due to its thyroid-stimulating activity (Nakabayashi et al., 2002). However, the thyrostimulin a-subunit (GPA2) is homologous but not identical to the common alpha subunit (GPA1 or a) in the other GpHs. With the discovery of GPA2 and GPB5 (thyrostimulin-b) homologs in invertebrates (including Drosophila melanogaster), it was proposed by Sudo, Kuwabara, Park, Hsu, and Hsueh (2005) that an ancestral heterodimeric GpH hormone existed before the divergence of vertebrates/invertebrates, and that a later gene duplication event in vertebrates produced thyrostimulin (GPA2 and GPB5) and GTH/TSH (GPA1 and LHb/FSHb/TSHb). This ancestry of GpH is supported by recent studies in which GPB5 (Tando & Kubokawa, 2009; Dos Santos et al., 2009) and GPA2 (Dos Santos et al., 2009) were identified from amphioxus, a basal chordate. Preliminary evidence shows only one glycoprotein a-subunit in lampreys that has higher similarity with mammalian GPA2 compared to the GPA1 (a) subunits (Sower, Freamat, & Kavanaugh, 2009). Despite extensive research using both molecular cloning and blasting of the lamprey genome, no evidence of a GPAa has been found; however, there is preliminary data for a GPB5 (Sower et al., 2009). Thus, it appears that lampreys may have ancestral GPA2, GTHb, and GPB5, but this will have to be confirmed by cloning. These authors’ current working hypothesis is that the GPA2 identified is the ancestral asubunit; after the gnathostomeeagnathan divergence, gene duplications produced the two a-subunits (GPA1 and GPA2) and four b-subunits (FSHb, LHb, TSHb, GPBb), and potentially lampreys may have two GpHs: GPA2/b and/or GPA2/GPB5 (Fig. 2) (Sower et al., 2009).

3.3.5. Glycoprotein hormone (GpH) receptors Gonadotropin and TSH hormone actions are mediated through a subfamily of GPCRs, namely the GpH receptors


(Combarnous, 1992). Known GpH-Rs share a number of unique features. They are composed of two functionally distinct modules of similar size, an extracellular N-terminal domain followed by a prototypical GPCR segment. The extracellular N-terminal domain is primarily responsible for high-affinity hormone binding and contains a central portion of nine Leu-rich repeat motifs, flanked by N- and C-terminal Cys-rich clusters. The C-terminal half of the receptor contains a transmembrane region with seven hydrophobic transmembrane a-helices, connected by intraand extracellular loops and an intracellular C-terminal domain (Grossmann, Weintraub, & Szkudlinski, 1997; Dufau, 1998; Ascoli, Fanelli, & Segaloff, 2002; Moyle et al., 2005). To date, approximately 79 GpH-Rs have been identified and described in 36 different species, mostly in mammals and also in three species of bird, two species of reptile, one amphibian, and ten species of fish (Hovergen Database). Until recently, there had been no GpH-Rs described in any species of agnathan. One functional GpH receptor (lGpH-R I) (Freamat, Kawauchi, Nozaki, & Sower, 2006) was identified from lamprey testis and a second functional GpH-R receptor (lGpH-R II) was identified and shown to be expressed mainly in thyroid tissue (Freamat & Sower, 2008). These authors hypothesize that lGpH-R I and lGpH-R II are the only members of the GpH receptor subfamily in lampreys (Freamat & Sower, 2008). They are descendants of the TSH receptor-like molecular ancestors of the GpH-Rs in gnathostomes and are likely the result of the genome duplication event hypothesized to have taken place before the divergence of the lamprey lineage (Sidow, 1996; Kuratani, Kuraku, & Murakami, 2002). The 719-amino-acid full-length cDNA encoding lGpH-R I is highly similar to and likely a homolog of the vertebrate GpH-Rs (including LH, FSH, and TSH receptors) (Freamat et al., 2006). The key motifs, sequence comparisons, and characteristics of the identified GpH-R reveal a mosaic of features common to all other classes of GpH-R in vertebrates. The lGpH-R I has been shown to activate the cAMP signaling system using hCG in transiently transfected COS7 cells. The highest expression of the receptor transcript has been demonstrated in the testes using reverse transcription polymerase chain reaction (PCR). The high expression of lGpH-R I in the testis and the high similarity with gnathostome gonadotropin hormone receptors suggest that lGpH-R I functions as a receptor for lamprey GTH hormones. The second GpH receptor (lGpH-R II) in the sea lamprey has 781 residues protein. IGPH-R II is approximately 43% identical compared to mammalian TSH-R and FSH-R representative amino acid sequences (Freamat & Sower, 2008). Similarly to these two classes of

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mammalian receptor, lGpH-R II is assembled from 10 exons. A synthetic ligand containing the lamprey GpH b chain tethered upstream of a mammalian a chain activates the lGpH-R II expressed in COS-7 cells, but to a lesser extent than lGpH-R I. The most obvious feature of the coding sequence of lGpH-R II is the presence of a long linker fragment called the ‘signaling specificity domain’ (SSD) or ‘hinge’ located between the Leu-rich domain (LRD) of the extracellular segment and the transmembrane domain. This is one of the longest linker fragments described in all vertebrate GpH receptors. This is in contrast with the similar region of the lGpH-R I, which is the shortest SSD/hinge segment among all vertebrate GpH-Rs (Freamat et al., 2006). Molecular phylogenetic analysis of vertebrate GpH-R protein

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sequences suggests a closer relationship between lGpH-R II and gnathostome TSH receptors. Therefore, at this point, a comparative perspective on this endocrine compartment in the lamprey relative to the well-established gnathostome paradigm suggests the involvement of one pituitary GpH (only b-subunits have been found) and two GpH-Rs as opposed to three or four dimeric hormones and three receptors in gnathostomes (Figure 10.2). This role of the GpH/GpH-R system in the lamprey has yet to be confirmed experimentally. This requires identification and characterization of a GpH achain homolog. The existence of a second GpH ligand in the lamprey with a distinct binding specificity to GpH-R I and II cannot be excluded, but, if it exists, it is likely even less similar to the gnathostome sequences than the one

FIGURE 10.2 Sower, Freamat, and Kavanaugh (2009) hypothesize that the hypothalamicepituitaryegonadal (HPG) and hypothalamicepituitaryethyroid (HPT) endocrine systems evolved from an ancestral, prevertebrate, exclusively neuroendocrine mechanism by gradual emergence of the components of a new control level (GpHs/GpH-Rs) concomitantly with the development of the corresponding anatomical structure (pituitary). The endocrine control of reproductive and thyroid functions in lampreys may reflect an intermediary stage on the evolutionary pathway to the highly specialized gnathostome HPG and HPT axes (Sower et al., 2009). CNS, central nervous system; GnRH, gonadotropinreleasing hormone; TRH, thyrotropin-releasing hormone; lGpH, lamprey glycoprotein hormone; lGpHR-I, lamprey glycoprotein hormone-receptorI; lGpHR-II, lamprey glycoprotien hormone-receptor-II; T3, triiodothyronine; T4, thyroxine; CRH, corticotropin-releasing hormone; TSH, thyrotropin or thyroid stimulating hormone; GTH1, gonadotropin 1; GTH2, gonadotropin 2; FSH, follicle-stimulating hormone; LH, luteinizing hormone; FSH-R, follicle-stimulating hormone receptor; TSH-R, thyroid stimulating hormone receptor; LHR, luteinizing hormone receptor. See color plate section.

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already described (Freamat & Sower, 2008). From the studies completed to date, these authors hypothesized that there is lower specificity of GTH and its receptor in agnathans and that, during coevolution of the ligand and its receptor in gnathostomes, there were increased specificities of interactions between each GpH (TSH, LH, and FSH) and its receptor (Figure 10.3).

3.3.6. Reproductive steroids The physiological role of gonadal sex steroids and their target sites in lampreys has mostly been studied in the sea lamprey, with a few studies in other species of lamprey. There are still many questions remaining as to the types of


steroid that are synthesized and their respective functions (Bryan, Scott, & Li, 2008). As cited in Bryan et al. (2008), much more research is needed to elucidate and understand whether lampreys use classical steroids and/or 15ahydroxylated steroids, as well as to identify receptors. In earlier studies, plasma E2 and P4 were measured as indicators of gonadal activity in sea lampreys under various physiological conditions. Estradiol (Katz, Dashow, & Epple, 1982; Fukayama & Takahashi, 1985; Sower, Plisetskaya, & Gorbman, 1985a) and P4 (Linville, Hanson, & Sower, 1987; Sower, 1989) have been associated with reproductive activity in both female and male sea or Japanese (Lampetra japonica) lampreys. In an earlier study, plasma E2 but not testosterone (T) was elevated in response

FIGURE 10.3 A proposed evolution of the glycoprotein hormone (GpH) subunits and glycoprotein hormone receptor (GpH-R) in vertebrates. (Bottom right) Freamat and Sower (2008) hypothesize that lGpH-R I and lGpH-R II are the only members of the GpH-R subfamily in lampreys. They are descendants of the thyrotropin receptor-like molecular ancestors of the GpH-Rs in gnathostomes and are likely the result of the genome duplication event hypothesized to have taken place before the divergence of lamprey lineage. (Bottom left) Ancestral glycoprotein subunits (GPA, a) and (GPB5, b) likely existed in a common ancestor of invertebrates and vertebrates (Sudo, Kuwabara, Park, Hsu, & Hsueh, 2005). Sower, Freamat, and Kavanaugh (2009) propose that an ancestral GpH gave rise to only one gonadotropin, and to the GpH family that gave rise to luteinizing hormone (LH), follicle-stimulating hormone (FSH), and thyroid-stimulating hormone (TSH) during the early evolution of gnathostomes. Lampreys may have one ancestral GPA2 and one, possibly two, beta subunits: GPb and/or GPB5 (unpublished data). The authors’ current working hypothesis is that the glycoprotein hormone subunit A2 identified is the ancestral a-subunit; after the gnathostomeeagnathan divergence, gene duplications produced the two a-subunits (GPA1 and GPA2) and three b-subunits (FSHb, LHb, and TSHb). Lampreys may have two GpHs: GPA2/b and/or GPA2/GPB5 (Sower et al., 2009). (Top) From the studies completed to date, Sower et al. (2009) hypothesize that there is lower specificity of gonadotropin (GTH) and its receptor in agnathans and that, during coevolution of the ligand and its receptor in gnathostomes, there were increased specificities of interactions between each GpH (TSH, LH, and FSH) and its receptor, depicted by the increase of shading from left to right. Thyrostimulin, another glycoprotein, is depicted by B5/A2 GpH. To date, thyrostimulin has only been shown to interact with TSH-R in gnathostomes. The question marks indicate the possible interactions of the subunits of the lamprey GpHs that are currently under investigation (Sower et al., 2009). LGR, Leucine-rich repeat-containing G-protein-coupled receptor. See color plate section.

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to a mammalian GnRH analog in male and female sea lampreys (Sower, Plisetskaya, & Gorbman, 1985b). In European river lampreys (Lampetra fluviatilis), plasma E2 was associated with vitellogenesis in females and spawning in males (Barannikova, Boev, Arshavskaya, & Dyubin, 1995; Mewes, Latz, Golla, & Fischer, 2002). These studies and the demonstrated absence of androgen receptors (Ho, Press, Liang, & Sower, 1987) in the lamprey testis suggested that T may not have a role during the final spermatogenetic phases in adult male lampreys. Both the ovary in brook lampreys (Lampetra aznandreal) (Belvedere & Colombo, 1983), and the testis in sea lampreys (Callard, Petro, & Ryan, 1980) have been shown to be capable of synthesizing E2. These data suggest that at least one of the classical steroidsdE2dhas a major role in aspects of reproduction in lampreys. The importance of E2 in lamprey reproduction is further supported by recent information on the cloning of steroid receptors. The first steroid receptorsdlamprey P4-like receptor (PR), the estrogen-like receptor (ER), and a corticoid-like receptordhave been identified (Thornton, 2001). Thornton (2001) proposed that the first steroid receptor in vertebrates was an ER, followed by a PR. This was based on identification and phylogenetic analysis of steroid receptors in basal vertebrates and reconstruction of the sequences and functional attributes of ancestral proteins. The androgen receptor (AR) was not identified and is proposed to have been created by a gene duplication after the lamprey lineage diverged from other vertebrates (Thornton, 2001). Specific regulation of physiological processes by androgens and corticoids are unique to gnathostomes and are proposed in this study to be relatively recent innovations that emerged after these duplications. Thus, from this study and the information stated above, estrogen regulation of reproductive maturation and function appears to be a major steroid control in both male and female lamprey reproduction. Therefore, T should be considered only a precursor for E2 synthesis. Recent studies have shown that lampreys produce and circulate 15a-hydroxylated steroids (Bryan et al., 2008). Kime and Rafter (1981) and Kime and Callard (1982) first found 15- hydroxylated compounds produced in the gonads of river and sea lampreys. Based on the effects of partial hypophysectomy and gonadectomy on plasma levels of various steroids in river lampreys (L. fluviatilis), Kime and Larsen (1987) later suggested that the true sex hormones responsible for development of secondary sex characteristics may be 15-hydroxylated derivatives of E2 and T. More recently, it was shown that steroids produced in vivo and in vitro coeluted with 15a-hydroxylated steroids on high performance liquid chromatography (HPLC) in all life phases of sea lampreys (Lowartz et al., 2003; Lowartz et al., 2004). This was verified by the identification of 15ahydroxytestosterone and 15a-hydroxyprogesterone as the

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major products of T and P4 in the lamprey testis using various chromatographic and immunological techniques as well as measurement of these steroids in plasma of male lampreys by specific radioimmunoassay (Bryan, Scott, Cerny, Seon Yun, & Li, 2003; Bryan, Scott, Cerny, Young, & Li, 2004). It will be important to identify specific receptors for these steroids to fully understand the functional nature of these unique steroids.

3.4. Circulating Hormones During Reproductive Cycles Lampreys are clearly seasonal and temperature-responsive in the timing of their anadromous migrations and reproduction. Many of these processes are coordinated by the neuroendocrine axis through a number of different hormones. As previously described, there are progressive changes in hormonal levels over time that occur in a changing endocrine environment (Sower, 2003). It has now been well-documented that an essential environmental cue to initiate metamorphosis in sea lampreys is the increase in water temperature in the spring (Holmes, Beamish, Seelye, & Youson, 1994; Holmes & Youson, 1994). As described by Youson (1997), circulating concentrations of the thyroid hormones, thyroxine (T4) and triiodothyronine (T3), drop dramatically at the onset of metamorphosis and do not appear to regulate lamprey metamorphosis as observed in other vertebrate metamorphoses, but these hormones are important in earlier developmental processes. As an indication of the complex simultaneous changes in lampreys undergoing metamorphosis, there is an increase of brain lamprey GnRH-I and -III that coincides with acceleration of gonadal maturation (Youson & Sower, 1991). Relatively little is known of circulating reproductive hormones coinciding with the reproductive maturation during the parasitic phase. However, in adult lampreys, there are seasonal correlations between changes in brain GnRH and gametogenic and steroidogenic activity of the gonads in adult male and female sea lampreys (Fahien & Sower, 1990; Bolduc & Sower, 1992). In sea lamprey females, lamprey GnRH-III is present in higher concentrations than lamprey GnRH-I during the final stages (as described in Section 3.3.2) of the reproductive season (MacIntyre, Chase, Tobet, & Sower, 1997). Lamprey GnRH-I concentrations do not change significantly during the reproductive season, whereas lamprey GnRH-III undergoes significant increases during the same period. These results suggest that lamprey GnRHIII may be the major form regulating reproductive processes in the female sea lamprey during the period of final reproductive maturation. In lampreys, reproductive activities include maturation, migration, and mobilization of fat stores. For example, as

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reviewed by Larsen and Dufour (1998), adult lampreys do not feed during the final reproductive period and there is significant mobilization of body tissue used to cover energy requirements for gonadal growth. It is proposed that a number of different endocrine systems could be involved in these various activities, including thyroid and reproductive hormones. Annual cycles in the gonads and thyroid gland and their interactions have been described for numerous species of vertebrate. Several studies of other fish species have indicated that sex steroids can influence thyroid activity (Sower, Sullivan, & Gorbman, 1984). Plasma T4 was significantly elevated in female sea lampreys following administration of a GnRH analog (Sower et al., 1985b). Not surprisingly, there are also coordinated changes in plasma T3 and T4 in males and females during the final maturational period. Plasma T3 shows a significant peak coinciding with a significant drop of T4 early in the final maturational period. Whether these thyroid hormones are coordinated or independently associated with lamprey reproduction is not known. However, it is evident that there are changes of hormones associated with the final maturational period and migration.

4. SUMMARY The phylogenetic relationship between the hagfish, the lamprey, and the jawed vertebrates is an unresolved issue (Kuraku, Hoshiyama, Katoh, Suga, & Miyata, 1999; Delarbre, Gallut, Barriel, Janvier, & Gachelin, 2002). In this chapter, agnathans are considered to be monophyletic in origin with the modern agnathans, which are classified into two groups, myxinoids (hagfish) and petromyzonids (lamprey), while the gnathostomes constitute all the other living vertebrates, including the bony and cartilaginous fishes and the tetrapods. It should be recognized, though, that the lampreys and hagfish diverged from each other hundreds of millions of years ago and have very different lifestyles, as described above. It is generally believed that two large-scale genome duplications (2R) occurred during the evolution of early vertebrates, although there is controversy on whether the 2R duplications occurred before the divergence of the hagfish and lampreys, or whether there was one round prior to the jawless vertebrates and one round after the divergence of the jawless vertebrates (Ohno, 1970; Holland, Garcia-Fernandez, Williams, & Sidow, 1994; Irvine et al., 2002; Larhammar, Lundin, & Hallbook, 2002; Fried, Prohaska, & Stadler, 2003; Vandepoele, De Vos, Taylor, Meyer, & Van de Peer, 2004; Kuraku, Kuraku, Meyer, & Kuratani, 2009). Further information on the evolution of vertebrate brain/pituitary hormones and their genes in lamprey and hagfish would contribute to the ongoing phylogenetic analysis that may


help in resolving the phylogenetic relationships between hagfishes, lampreys, and jawed vertebrates. There are marked similarities yet significant differences in the neuroendocrine systems of hagfishes and lampreys, which in part may reflect the origin and ancestry of these extant agnathans. The endocrine control of reproductive and thyroid functions in lampreys reflects an intermediary stage on the evolutionary pathway to the highly specialized gnathostome HPG and hypothalamicepituitaryethyroid (HPT) axes. The information on GTHs and GpH in lampreys have established that, similar to all other vertebrates, lampreys (freshwater dwellers or anadromous) have a HPG axis and that there is a high conservation of the mechanisms of GnRH action. On the other hand, in hagfish that live in a deep marine environment, there seems to be a lack of, or poor regulation of, reproduction by hypothalamicepituitary peptides (Sower & Gorbman, 1999). The extent to which the modern hagfishes and lampreys represent or reflect the ancient forms from which they evolved 500 million years ago is a question that still concerns evolutionary biologists and that is far from being answered. The ancient origin of the lamprey and hagfish lineages and the relative conservation of ancestral features suggest that the study of these two distinct ancient lineages may offer the premise for a more accurate estimation of how the genome, proteome, and signaling networks of the common ancestor of vertebrates may have looked.

ABBREVIATIONS 2R ACTH AR CG COS7 C-terminal E2 ER FSH GnRH GnRH-R GPA2 GPB5 GPCR GpH GpH-R GTH HPG HPLC HPT lGnRH I lGnRH II lGnRH III lGpH-R 1

Two whole rainds of genome duplication Corticotropin Androgen receptor Chorionic gonadotropin Mammalion cell line Carboxy-terminal Estradiol Estrogen-like receptor Follicle-stimulating hormone Gonadotropin-releasing hormone Gonadotropin-releasing hormone receptor Thyrostimulin-a Thyrostimulin-b G-protein-coupled receptor Glycoprotein hormone Glycoprotein hormone receptor Gonadotropin Hypothalamicepituitaryegonadal High performance liquid chromatography Hypothalamicepituitaryethyroid Lamprey GnRH I Lamprey GnRH II Lamprey GnRH III Lamprey GpH receptor 1

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lGpH-R 2 IP3 ir LH LRD P4 PCR PR RT-PCR SSD T T3 T4 TSH


Lamprey GpH receptor 2 Inositol triphosphate Immunoreactive Luteinizing hormone Leu-rich domain Progesterone Polymerase chain reaction Progesterone-like receptor Reverse transcription-polymerase chain reaction Signaling specificity domain Testosterone Triiodothyronine Thyroxine Thyrotropin or thyroid-stimulating hormone

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Chapter 11

Hormones and Reproduction in Chondrichthyan Fishes Karen P. Maruska* and James Gelsleichtery *

Stanford University, Stanford, CA, USA, y University of North Florida, Jacksonville, FL, USA

SUMMARY Chondrichthyans represent a diverse and successful group of fishes that occupy a critical position in the evolution of vertebrate animals. The evolutionary success of these fishes is partly attributed to their many reproductive adaptations, and an understanding of reproductive endocrinology in this group can provide insights into hormonal function in all vertebrates. This chapter summarizes the current knowledge of the role of hormones in the reproductive physiology of chondrichthyan fishes, and identifies important areas for future research. The roles of peptide and steroid hormones in both males and females are discussed in relation to the brainepituitaryegonadal axis, regulation of the reproductive tract and gametogenesis, sexual maturation, mating behavior, and environmental influences on reproduction. Recent studies on chondrichthyan endocrinology have provided important information on how hormones regulate reproduction in this diverse group of fishes, and demonstrate that many regulatory mechanisms are conserved through vertebrate evolution.

1. REPRODUCTION IN CHONDRICHTHYAN FISHES The cartilaginous fishes (class Chondrichthyes) include the well-known subclass Elasmobranchii (sharks, skates, and rays) and the smaller, less understood, Holocephali (chimaeras, ratfishes, and rabbitfishes). Chondrichthyan fishes have a long evolutionary history that goes back over 400 million years. Part of their evolutionary success is attributed to their diverse reproductive modes and adaptations related to reproductive physiology. For example, all chondrichthyan fishes have internal fertilization, where the male mates with the female and uses one of his paired claspers (intromittent copulatory organs) to transfer sperm to her reproductive tract for egg fertilization. This internal fertilization, coupled with efficient maternal nourishment of embryos and the birth of large, well-developed offspring, contributes to the enormous success of these fishes. Internal Hormones and Reproduction of Vertebrates, Volume 1dFishes Copyright Ó 2011 Elsevier Inc. All rights reserved.

fertilization and the many complex reproductive modes have led to the notion that cartilaginous fishes are reproductively more similar to reptiles, birds, and mammals than to other fishes. This great diversity in reproductive adaptations also requires a complex hormonal system that regulates all aspects of reproduction from sexual maturity to mating behavior and seasonal changes in gonadal physiology (Figure 11.1).

1.1. Reproductive Modes Chondrichthyan fishes can be separated into two main groups, based on reproductive mode: oviparous (egglaying) and viviparous (live-bearing), the latter of which can be further divided into placental and aplacental forms. Oviparous species retain their eggs for varying amounts of time, enclose them in a protective egg case, and then deposit them on some substrate for development. Oviparity is the reproductive mode found in all of the Holocephali and the skates (Elasmobranchii; Rajiformes), and occurs in several families of sharks (e.g., Orectolobidae, Scyliorhinidae, Heterodontidae) (Wourms, Grove, & Lombardi, 1988). Oviparity is also thought to be the ancestral condition in chondrichthyan fishes, from which viviparity arose independently ~9e10 times (Wourms, 1977; Dulvy & Reynolds, 1997). However, a more recent analysis argues that viviparity is the ancestral state in chondrichthyans, and that oviparity then evolved to increase fecundity in smallbodied species that have limited coelomic space for numerous live offspring (Musick & Ellis, 2005). Viviparity is characteristic of all rays and occurs in about 70% of all shark species. Viviparous species retain their embryos in the uterus until they are fully developed and then give birth to live young that are miniature versions of the adults. Viviparity can be divided into placental and aplacental varieties, depending on whether a placental connection exists between the mother and embryo. In placental species, the embryo is initially nourished by 209

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FIGURE 11.1 Summary of the proposed mechanism for hormonal regulation of chondrichthyan reproduction. Male reproductive tract is on the left, and female on the right. Open circle, ultimobranchial gland; closed circle, thyroid gland. CT, calcitonin; DHT, dihydrotestosterone; E2, 17bestradiol; GnRH, gonadotropin-releasing hormone; GTH, gonadotropins; P4, progesterone; Rlx, relaxin; T, testosterone; T3, triiodothyronine; T4, thyroxine. See text for additional details. Reproduced from Gelsleichter (2004), with permission.

a yolk sac, which, when depleted, elongates and forms a highly vascularized connection with the uterine wall of the mother to form a yolk sac placenta. Nutrients are then delivered directly from the bloodstream of the mother to the developing embryo. Placental elasmobranchs generally have long gestation periods, relatively low numbers of offspring (a few to over a hundred depending on the species), increased maternal protection during embryonic development, and greater survival at birth due to the large size of fully developed young. At birth, the umbilical cord (former yolk stalk) breaks off and leaves a small scar on the body between the pectoral fins of the newborn offspring.


The status of this umbilical scar (from fresh/open to fully healed) is often used by researchers to determine the relative age of newborn pups as well as the seasonal timing of parturition (birth). Aplacental viviparity can be separated into three functional groups based on how the embryo receives its nutrition: (1) yolk dependency: embryos depend solely on the yolk deposited in the egg at ovulation, with no supplemental nourishment from the mother. This type is found in Squaliformes, Hexanchiformes, Squatiniformes, and some Carcharhiniformes, Orectolobiformes, and rays. (2) Intrauterine cannibalism: the most common of these forms, oophagy (egg-eating), occurs when embryos develop initially on yolk reserves and then begin to ingest a supply of unfertilized eggs in the uterus with precocial teeth. This type is found primarily in the lamnoid sharks (e.g., mako, thresher, great white) (Gilmore, Putz, & Dodrill, 2005), but may also exist in Carcharhinidae and Orectolobidae families. In the sandtiger shark Carcharius taurus, a more extreme form called embryophagy occurs when the largest and strongest embryo(s) develops quickly and then consumes the other developing young within the uterus. (3) Placental analogs: embryos are nourished by regions of the uterine epithelium called trophonemata that secrete a ‘uterine milk’ or ‘histotroph,’ a process known as uterolactation. This type is prevalent in the majority of rays (Myliobatiformes). For a more detailed description of the evolution and diversity of reproductive modes found in chondrichthyan fishes, the reader is directed to the following references: Wourms (1977); Wourms et al. (1988); Wourms and Demski (1993); Carrier, Pratt, and Castro (2004); Gilmore et al. (2005); Hamlett, Kormanik, Storrie, Stevens, and Walker (2005); Musick and Ellis (2005).

1.2. Reproductive Cycles Annual reproductive cycles in chondrichthyan fishes are complex and diverse, but most species can be grouped into one of three different types based on the ovarian cycle and gestation period in females (Hamlett & Koob, 1999; Koob & Callard, 1999). Continuous breeders (many viviparous sharks, some batoids) are reproductively active year-round with pregnancy lasting almost a full year, and mating, pregnancy, and parturition are generally coupled with environmental factors and synchronized within the population. Seasonal breeders (most ray species, some viviparous sharks) are reproductively active for only a portion of the annual cycle, with pregnancy lasting several months and the remainder of the year spent nonpregnant. Similar to continuous breeders, mating, pregnancy, and parturition are correlated with environmental cues and occur at approximately the same time each year to synchronize the population. Punctuated breeders (some viviparous sharks and

Chapter | 11

Hormones and Reproduction in Chondrichthyan Fishes

rays, some oviparous species) are often pregnant for about a full year, but spend one or more intervening years in a nonpregnant state. For example, in blue (Prionace glauca) and sandbar (Carcharhinus plumbeus) sharks, females are pregnant for a full year but only deliver young every two years, spending the intervening year nonpregnant. There are also cases of 2e3.5 year pregnancies in some elasmobranchs (e.g., Squalus acanthias, Chlamydoselachus anguineus) and new studies continue to describe diverse reproductive cycles in chondrichthyans.

1.3. Mating and Reproductive Behaviors Direct observations of courtship and mating behaviors in chondrichthyans are rare, especially in the wild, but the available descriptions reveal a complex suite of reproductive behaviors in this group of fishes including dominance hierarchies, pair and group mating behaviors, sexual segregation, and cooperative breeding (Pratt & Carrier, 2001; Carrier et al., 2004; Pratt & Carrier, 2005). Precopulatory behaviors (e.g., following, parallel swimming, biting, female avoidance and acceptance, clasper flexion) may be brief or prolonged, but ultimately culminate in the male grasping the female on the fin or body so that clasper insertion and internal fertilization can occur. This oral grasping often leaves mating scars on the female, which can be used as an indicator of active courtship and/or mating season (Kajiura, Sebastian, & Tricas, 2000). Once the female accepts (often becoming immobile and flaring or cupping her pelvic fins) and the male has an adequate grip, a single clasper is rotated forward, inserted, and often anchored in the female by opening of the terminal cartilages that bear a hook or spur in some species, and semen is then transferred into the female’s reproductive tract. Copulation can be brief (seconds to minutes), or may last for several hours, as observed in some benthic skate species maintained in captive settings (Luer & Gilbert, 1985). Females of several elasmobranch species can also store sperm (for weeks to months) in the reproductive tract for later fertilization (Pratt, 1993; Hamlett, Knight, Pereira, Steele, & Sever, 2005). Females may mate with several males, and multiple paternity has been documented in several elasmobranch species, possibly to maintain genetic diversity in animals that produce only a few broods during their lifetime (Chapman, Corcoran, Harvey, Malan, & Shivji, 2003; Carrier et al., 2004; Feldheim, Gruber, & Ashley, 2004; Heist, 2004; Pratt & Carrier, 2005; Daly-Engel, Grubbs, Holland, Toonen, & Bowen, 2006; Chevolot, Ellis, Rijnsdorp, Stam, & Olsen, 2007; Portnoy, Piercy, Musick, Burgess, & Graves, 2007). However, high rates of single male paternity also occur in certain species, such as the bonnethead shark, suggesting that pre- and/or postcopulatory factors that influence the mating systems of these animals may be diverse (Chapman, Prodohl,

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Gelsleichter, Manire, & Shivji, 2004). In contrast to most other vertebrate taxa, parental care has not yet been described in chondrichthyan fishes.

2. THE HYPOTHALAMICePITUITARYe GONADAL (HPG) AXIS As in all vertebrates, reproduction in chondrichthyan fishes is regulated by the hypothalamicepituitaryegonadal (HPG) axis. External cues from the environment such as photoperiod and temperature, or internal physiological cues associated with sexual maturation (or puberty), initiate the endocrine cascade that begins with activation of the gonadotropin-releasing hormone (GnRH) neurons in the brain. Gonadotropin-releasing hormone then stimulates production and release of the gonadotropin (GTH) hormones from the pituitary into the bloodstream, where they travel to the gonads (ovary and testis) to promote gametogenesis and steroidogenesis (Figure 11.1). Gonadal steroids are also responsible for further regulating gamete production (via autocrine and paracrine mechanisms in the gonads and feedback mechanisms to the brain and pituitary), and, presumably, secondary sex characteristics, sensory function, and reproductive behavior. While the basic structure and function of the HPG axis is conserved among vertebrates, there are several distinct differences found in the chondrichthyan fishes compared to both higher and lower taxa that are discussed below.

2.1. Gonadotropin-releasing Hormone (GnRH) Gonadotropin-releasing hormone is represented by a family of decapeptides produced in the brain of all vertebrates and is a critical regulator of reproduction (see Chapter 2, this volume). In chondrichthyan fishes, distinct populations of GnRH neurons are found in the basal forebrain or preoptic area (POA) (GnRH1), the terminal nerve (TN) ganglia, and the midbrain tegmentum (GnRH2) (Figure 11.2) (Sherwood & Lovejoy, 1993; Demski, Beaver, Sudberry, & Custis, 1997). Each of these regions expresses a GnRH variant with a slightly different amino acid sequence. In all chondrichthyan fishes studied to date, multiple forms (i.e., as many as seven) of GnRH are found in the brain of the same species, and in some species multiple forms are localized to the same brain region (Powell, Millar, & King, 1986; Lovejoy et al., 1991; 1992; D’Antonio et al., 1995; Demski et al., 1997; Forlano, Maruska, Sower, King, & Tricas, 2000; Masini, Prato, Vacchi, & Uva, 2008). Despite their important taxonomic position and diversity of reproductive modes, there are no published molecular studies on the sequence or gene structure of chondrichthyan GnRH peptides or GnRH receptors. As a result, recent analyses on

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FIGURE 11.2 Distribution of gonadotropin-releasing hormone (GnRH)-immunoreactive (ir) somata and fibers in the elasmobranch brain. Diagrammatic parasagittal representations show the distribution of forebrain (a) and midbrain (b) GnRH-ir somata (dots) and fibers (lines) in the brain of the Atlantic stingray (Dasyatis sabina). (c) Representative monopolar GnRH-ir cell in the preoptic area of D. sabina. (d) Representative sagittal section showing the large GnRH-ir cell group in the midbrain of D. sabina. (e) Schematic drawing of the terminal nerve (TN) and ganglia on the dorsal forebrain of D. sabina. Inset shows a monopolar GnRH-ir neuron (arrow) from the white body (WB) of the TN. (f) Summary schematic diagram illustrates the GnRH-ir fiber pathways (arrows) in the elasmobranch brain. Hatched area represents the midbrain GnRH-ir cell group. Numbers 1e3 represent the positions of terminal nerve entry in different species: (1) Urobatis (Urolophus) halleri and Platyrhinoidis triseriata; (2) Squalus acanthias and Triakis semifasciata; and (3) Triakis semifasciata. 3V, third ventricle; CE, corpus cerebellum; DON, dorsal octaval nucleus; G2, ganglion 2 of TN; G3, ganglion 3 of TN; HA, habenula; HP, hypothalamus; LP, lateral pallium; M, medulla; ML, median lobe of pituitary; NIL, neurointermediate lobe of pituitary; OB, olfactory bulb; OC, optic chiasm; OE, olfactory epithelium; OLT, olfactory tract; PC, posterior commissure; pi, pituitary; POA, preoptic area; RL, rostral lobe of pituitary; se, septal area; SV, saccus vasculosus; T, tectum; TEG, tegmentum; TEL, telencephalon; v, ventricle. Scale bars ¼ 0.5 cm (a, b); 20 mm (c, e inset); 200 mm (d); 1 cm (e). (aee) Modified from Forlano et al. (2000), with permission. (f) Modified from Wright and Demski (1993), with permission.

the molecular evolution of GnRH and its receptor have not included any chondrichthyan representatives (Guilgur, Moncaut, Canario, & Somoza, 2006; Flanagan et al., 2007; Chen & Fernald, 2008; Kavanaugh, Nozaki, & Sower, 2008; Okubo & Nagahama, 2008). Until the localization of distinct GnRH molecular variants within the TN ganglia, POA, hypothalamus, and midbrain tegmentum are determined in chondrichthyans, it will be difficult to fully appreciate the function and evolutionary relationships of these different cell groups. Gonadotropin-releasing hormone neurons found in the forebrain region (e.g., ventral telencephalon, POA,

hypothalamus) of vertebrates generally project to the pituitary and are responsible for GTH production and release (i.e., the releasing form; GnRH1). Gonadotropinreleasing hormone neurons in the POA of chondrichthyan fishes do project to the rostral, median, and neurointermediate lobes of the pituitary (Demski et al., 1997; Forlano et al., 2000), but do not reach the isolated ventral lobe that contains the most GTH activity (see Section 2.2). In male Atlantic stingrays (Dasyatis sabina), the number of forebrain GnRH neurons varies with the seasonal reproductive cycle, consistent with a role in GTH control and gonadal recrudescence (Forlano et al., 2000). However,

Chapter | 11


chondrichthyans differ from most vertebrates in that GnRH is hypothesized to reach the ventral lobe via the general circulation, and there is some evidence for GnRH release to the bloodstream and cerebrospinal fluid from all three GnRH cell groups (Demski et al., 1997). Consequently, it remains unclear whether reproductive competence is regulated by a single (i.e., GnRH1) or multiple GnRH forms released by one or possibly several GnRH cell groups. The TN is a ganglionated cranial nerve that is distinct and separate from the olfactory nerve in chondrichthyans, and is another major source of GnRH in these fishes (Demski, Fields, Bullock, Schriebman, & Margolis-Nunno, 1987; Demski, 1993; White & Meredith, 1995; Demski et al., 1997; Forlano et al., 2000). There is some evidence that GnRH from the TN functions both in reproductive competence as a releasing factor (i.e., gonadal recrudescence and steroidogenesis) and in the modulation of sensory-mediated reproductive behaviors. Gonadotropinreleasing hormone-immunoreactive (ir) fibers are closely associated with both cerebral vasculature and ventricles (Demski & Fields, 1988; Demski et al., 1997), which provides a potential route for TN control of reproduction via GnRH release to the systemic circulation. Further, electrical stimulation of the TN in the Atlantic stingray results in increased GnRH levels in the cerebrospinal fluid, which provides support for GnRH access to other brain regions via an intraventricular route (Moeller & Meredith, 1998). Direct projections from the TN GnRH neurons to both the retina (Demski et al., 1997) and olfactory regions (e.g., olfactory bulb, and region between the olfactory bulb and olfactory epithelium) (Demski et al., 1997; Forlano et al., 2000) also suggest a role in the integration of visual and olfactory (i.e., pheromonal) cues with seasonal reproductive behaviors (Demski & Northcutt, 1983; Demski, 1991). As stated above, however, without an understanding of which specific GnRH variant(s) is found in the TN and whether this same form is present in the blood and CSF and has physiological effects on the gonads, the function of TN GnRH in chondrichthyans remains unclear. The GnRH neurons in the elasmobranch midbrain are an order of magnitude more numerous than those found in bony fishes (hundreds vs. tens), and have been shown to contain the evolutionarily conserved GnRH2 variant by both immunocytochemistry and radioimmunoassay (Figure 11.2) (Forlano et al., 2000). Gonadotropinreleasing hormone-ir axons from this midbrain group project to regions of visual, electrosensory, and mechanosensory processing in both the midbrain and hindbrain, as well as to motor neurons in the spinal cord that may regulate clasper movements (Forlano et al., 2000). Thus, it is hypothesized that midbrain GnRH influences reproductive success by modulating the sensitivity of the visual, electrosensory, somatosensory, and lateral line systems for

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mate detection and courtship behaviors, as well as regulating motor aspects of copulation including clasper movements. However, this neuromodulatory role for GnRH2 requires experimental confirmation.

2.2. Pituitary Structure and Gonadotropins (GTHs) As previously mentioned, the pituitary gland in elasmobranchs differs from that of bony fishes and higher vertebrate taxa because there is no direct neural or portal blood connection to deliver GnRH from the brain to the ventral lobe of the pituitary where the GTHs are produced (Figure 11.3) (J. Dodd, M. Dodd, & Duggan, 1983). Holocephalans, on the other hand, lack a ventral lobe and do have neural and vascular connections to the pituitary

FIGURE 11.3 Diagrammatic sagittal section of the hypothalamice pituitary gland vascular and neurosecretory connections in Holocephali and Elasmobranchii. In the species examined to date, direct vascular or neurosecretory connections to the ventral pituitary lobe appear to be absent in elasmobranchs. Holocephalans lack the ventral lobe entirely, but possess a buccal lobe (or rachendachhypophyse) (not shown) that is separated from the rest of the pituitary in mature adults. Dotted circles, neurosecretory cells of the nucleus preopticus; dotted lines, neurosecretory axons with axon terminals marked at their endings; filled circles, prehypophyseal plexus; open circles, intrahypophyseal capillaries; large interrupted arrows, arteries; large solid arrows, veins; thin arrows, portal veins. Rostral is to the left. rPD, rostral zone of pars distalis; pPD, proximal zone of pars distalis; NIL, neurointermediate lobe; aME, anterior median eminence; pME, posterior median eminence; SV, saccus vasculosus; VL, ventral lobe. Modified from Jasinski (1969), with permission.

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(Jasinski, 1969) (Figure 11.3). However, holocephalans also possess a buccal pituitary lobe, or ‘rachendachhypophyse,’ that in mature adults is completely separated from the rest of the pituitary but contains strong GTH activity (J. Dodd & M. Dodd, 1985). A capillary system does exist in elasmobranchs to join the hypothalamus with the rostral, median, and neurointermediate lobes of the pituitary, and in some species these lobes are also innervated by GnRH axons (Demski et al., 1997; Forlano et al., 2000), but a direct connection to the anatomically separate ventral lobe (elasmobranchs) or buccal lobe (holocephalans) has not been demonstrated (Dodd, 1975; Dodd et al., 1983; J. Dodd & M. Dodd, 1985). Instead, GnRH is hypothesized to reach the elasmobranch ventral lobe via an intraventricular route or the general circulation (King et al., 1992; Sherwood & Lovejoy, 1993; Demski et al., 1997; Moeller & Meredith, 1998). This is supported by the detection of both GnRH and GnRH-binding proteins in the cerebrospinal fluid and peripheral blood of many elasmobranchs (Powell et al., 1986; King et al., 1992; Pierantoni, D’Antonio, & Fasano, 1993; Sherwood & Lovejoy, 1993; D’Antonio et al., 1995; Moeller & Meredith, 1998). The ventral lobe is vascularized from the systemic circulation by the internal carotid artery, which forms a vascular bed (Dodd et al., 1983; Honma, Toda, & Chiba, 1987). This GnRH presence in the bloodstream at levels equivalent to the mammalian hypothalamoehypophysial portal system (Sherwood & Lovejoy, 1993), the fact that exogenous administration of GnRH analogs also influences gonadal steroidogenesis (Jenkins & Dodd, 1980; Fasano et al., 1989; Callard, Fileti, & Koob, 1993), and the result that ventral lobectomy only partially impairs gametogenesis and steroidogenesis (Dobson & Dodd, 1977a; 1977b; Sumpter, Jenkins, & Dodd, 1978b) also raise the possibility of direct actions of GnRH on the gonads in elasmobranchs. Gonadotropinreleasing hormone receptors have been detected in the gonads of bony fishes and other taxa, and therefore this putative direct hypothalamicegonadal connection that circumvents the pituitary gland certainly deserves future attention and experimentation. The presence of GTHs in the ventral lobe of the elasmobranch pituitary has been demonstrated by immunocytochemistry (Mellinger & DuBois, 1973) and radioimmunoassay (Scanes, Dobson, Follett, & Dodd, 1972), and several studies show that extracts of the ventral lobe can stimulate steroidogenesis in testicular cell culture (Lance & Callard, 1978; Sumpter, Jenkins, Duggan, & Dodd, 1980; Sourdaine, Garnier, & Jegou, 1990) as well as in vivo in both male and female elasmobranchs (Sumpter et al., 1978b ; Callard & Pasmanik, 1987). Extracts from the ventral lobe showed 10e100 times more GTH activity compared to intermediate and median pituitary lobe extracts, as measured by testosterone (T) production from dispersed reptilian testicular cells (Lance & Callard, 1978).


Further, hypophysectomy, or removal of the ventral lobe, alone causes partial regression of the testes and reduced androgen concentrations in some male elasmobranchs (Dodd, Evennett, & Goddard, 1960; Dobson & Dodd, 1977a; Sumpter, Follett, Jenkins, & Dodd, 1978; Fasano et al., 1989) and follicular atresia and impaired oviposition in female Scyliorhinus canicula (Norris, 1997). However, these removal experiments did not completely suppress gametogenesis or steroidogenesis and the responses may depend on reproductive stage and/or environmental stimuli (Dobson & Dodd, 1977c), a finding consistent with seasonal variation in pituitary GTHs. Further, it is important to note that the relative role of the ventral lobe in GTH release, and the regulation of gonadal steroidogenesis either via direct GnRH action or the traditional HPG axis, may be very different among diverse chondrichthyan species, especially those with different reproductive modes. For example, the majority of work on ventral lobe function is based on the dogfish S. canicula (Dodd, 1975; Dobson & Dodd, 1977a). However, in skates such as Leucoraja (Raja) erinacea, the ventral lobe is less developed and, although GnRH directly stimulates steroid synthesis in follicular cells of females in vitro, extracts of the ventral pituitary lobe do not (Callard et al., 1993; Demski et al., 1997). These differences in pituitary structure and function among the limited species examined thus far highlight that future comparative work on pituitary morphology, GTH localization and function, and the role of direct GnRH action in the gonads among sharks, skates, rays, and chimaeras is clearly needed. The GTHs, follicle stimulating hormone (FSH) and luteinizing hormone (LH), are both heterodimeric glycoproteins with identical a-subunits and distinct b-subunits. In most vertebrates, FSH targets the Sertoli cells of the testis and the follicular and granulosa cells of the ovary to stimulate gametogenesis. On the other hand, LH targets the interstitial Leydig cells of the testes and the granulosa and thecal cells of the ovaries and primarily functions to regulate steroidogenesis and final gamete maturation and release (spermiation and ovulation). One a-subunit and two distinct b-subunits have been cloned and sequenced from the ventral pituitary lobe of the dogfish S. canicula and the latter subunits are orthologs of FSH and LH found in bony fishes and tetrapods (Querat, Tonnerre-Doncarli, Genies, & Salmon, 2001). Thus, it seems plausible that the duality of the GTHs occurred prior to the Chondrichthyes. Experimental studies in male dogfish also indicate that GTHs are important for the transition from spermatogonia to spermatocytes, and that GTH levels fluctuate seasonally with the breeding cycle (Dobson & Dodd, 1977a; 1977b; 1977c). However, the differential distribution of FSH and LH receptors in the chondrichthyan gonad is unknown and evidence of discrete functions on steroidogenesis and gametogenesis is lacking.

Chapter | 11


3. HORMONAL REGULATION IN FEMALES 3.1. Structure and Function of the Female Reproductive Tract The general structure and morphology of the chondrichthyan ovary and other components of the reproductive tract have been reviewed in detail elsewhere (Hamlett & Koob, 1999; Carrier et al., 2004; Callard, St. George, & Koob, 2005; Hamlett et al., 2005a; Lutton, St. George, Murrin, Fileti, & Callard, 2005) and will only be briefly summarized here. The female chondrichthyan reproductive system consists of paired or single ovaries and the oviducts. Each oviduct is differentiated into an ostium, anterior oviduct, oviducal gland (also known as the shell or nidamental gland), isthmus, uterus, cervix, and the common urogenital sinus. In elasmobranchs, at least two different ovarian types are recognized based on the relationship between the ovary and the epigonal organ, a lymphomyeloid organ associated with the gonad in both sexes (Figure 11.4) (Pratt, 1988). ‘Internal’ ovaries, found in lamnoid sharks, have a germinal epithelium encapsulated within the epigonal organ and produce numerous small ova. In contrast, ‘external’ ovaries lie on the distal surface of the epigonal organ, produce fewer but larger eggs, and are found in many different elasmobranch species. Elasmobranch ova are large relative to bony fishes, and yolk accumulation is a slow process taking several months to a year in most species.

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Eggs released from an ovary are collected by the ostium (single or paired) and travel through the anterior oviduct to the oviducal gland, where they become fertilized, are surrounded by egg jelly, and are encapsulated by either rigid (in oviparous species) or more pliable (in viviparous species) egg capsules. These oviducal glands are unique to chondrichthyan fishes and all consist of morphologically distinct zones (i.e., the club, papillary, baffle, and terminal zones), which appear to perform different functions in the overall process of egg fertilization and encapsulation (Hamlett et al., 2005a). Oviducal glands vary tremendously among species, but their size and structural complexity correlates with reproductive mode, being larger and more complex in oviparous species that produce hard or leathery egg cases designed for external embryonic development and protection. In addition to egg jelly and tertiary egg envelope formation, oviducal glands in some species also function to store sperm within the female tract for later fertilization (Hamlett et al., 2005a). Following fertilization and encapsulation, eggs are transported from the oviducal gland to the uterus via a morphologically distinct component of the reproductive tract known as the isthmus. This region undergoes structural modifications associated with the reproductive cycle that first allow it to permit transit of fragile ova into the uterus, and later reduce its compliance to make the uterus an isolated compartment where modification of egg

FIGURE 11.4 Schematic cross-section representations of the different forms of elasmobranch testes (left panel: diametric, radial, compound) and ovaries (right panel: ‘external,’ ‘internal’). Left illustrations in each panel are diagrammatic with size of follicles (spermatocysts) exaggerated to show development. Arrows on right illustrations in each panel indicate paths of follicle or spermatocyst development (testis), and oocyte travel and release (ovary). Modified from Pratt (1988), with permission.

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capsules or embryonic development can take place. In oviparous species, the uterus holds the egg capsule during polymerization of egg capsule materials and tanning; a process collectively known as sclerotization. In viviparous species, the uterus houses the developing embryos until parturition occurs; a period during which it functions to regulate conditions in the intrauterine environment, supply oxygen for respiratory demands, and provide nutrients and waste disposal for growing young (Hamlett & Koob, 1999). This can involve a number of diverse morphological modifications to the uterine structure, such as significant enlargements in size; compartmentalization of the uterus to separate individual embryos; extensive folding and vascularization of the uterine epithelium to oxygenate uterine fluids or, in some species, produce various forms of matrotrophic nourishment (e.g., histotroph); and/or in placental viviparous species form often elaborate placental connections between pregnant females and offspring. Even in species in which embryos derive no additional nourishment beyond that supplied in yolk reserves, proper uterine function is critical to the wellbeing and survival of developing young. An often-cited example of such actions occurs in the aplacental viviparous shark S. acanthias, in which contractions of the myometrium serve to periodically flush seawater into the uterus, presumably to maintain electrolyte balance and remove embryonic wastes (Kormanik, 1993). During oviposition or parturition in oviparous and viviparous species, respectively, eggs or full-term young are released to the external environment through increased compliance of the previously rigid and inextensible cervix. In placental species, distinct scarring of the uterus occurs as a result of uterine compartmentalization and placentation, which then undergoes repair in preparation for the subsequent breeding season (Hamlett, Musick, Hysell, & Sever, 2002).

3.2. Steroidogenesis, Steroidogenic Enzymes, and Steroid Receptors As in other vertebrate groups, sex steroids including 17bestradiol (E2), T, and progesterone (P4) are produced by various cell types (i.e., granulosa cells, thecal cells, and corpora luteal tissue) in the ovary of female elasmobranchs (Figure 11.5). As previously discussed, this is likely regulated by pituitary GTHs; a premise supported by the reduction in steroidogenesis observed in certain elasmobranch species (i.e., the skate L. erinacea) following ventral lobectomy. Additionally, as illustrated by the stimulatory effects of GnRH on in-vitro and in-vivo gonadal steroidogenesis in L. erinacea and S. canicula, respectively, it is possible that the hypothalamus also may play a role in regulating ovarian steroid production in this group (Jenkins &


FIGURE 11.5 Tissue source and profiles of circulating 17b-estradiol (E2), testosterone (T), and progesterone (P4) titers during the ovulatory cycle of the female oviparous skate Leucoraja (Raja) erinacea. The principal source of each steroid is indicated by the solid lines in the upper panel, while additional sources are indicated by dashed lines. 17b-estradiol and T titers are elevated during the follicular phase of each ovulatory cycle, whereas P4 is elevated for only a brief period during the preovulatory phase and is low during ovulation, egg encapsulation, and oviposition. Modified from Koob and Callard (1999), with permission.

Dodd, 1980; Lutton et al., 2005). Lastly, as implied by its intimate association with the gonads in both sexes, the epigonal organ also appears to contribute to the regulation of gonadal steroidogenesis in elasmobranchs (Lutton & Callard, 2007; 2008a; 2008b). For example, recent studies have demonstrated that epigonal cells and conditioned media can inhibit in-vitro production of E2 and T in the skate ovary (Lutton et al., 2005), and, since blood flow from the gonadal artery first enters the epigonal organ and then perfuses the ovarian follicles, epigonal organ-secreted factors could directly influence follicular steroid production (Lutton & Callard, 2008b). Further, circulating gonadal steroids also may produce either stimulatory or inhibitory changes in leukocyte proliferation within epigonal tissue depending on physiological state and/or environmental cues (Lutton & Callard, 2008a), suggesting a functional interaction between reproductive and immune systems. As demonstrated for both the skate L. erinacea and the shark S. acanthias, E2 and T primarily are produced by both granulosa cells and theca cells of ovarian follicles (Figure 11.5) (Tsang & Callard, 1987b; Fileti & Callard, 1990; Tsang & Callard, 1992; Callard et al., 1993). However, the contribution of granulosa cells to E2 synthesis

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appears to be much greater than that of theca cells in at least some elasmobranchs. This is supported by in-vitro studies on steroidogenesis in granulosa cell and theca cell isolates from the skate ovary, which have demonstrated far greater increases in E2 production in the former in response to hormonal stimulation by GnRH (Lutton et al., 2005). In comparison to that for E2, T production by these two cell types is believed to be more equivalent. Although the follicles are generally considered to be the primary source of T in females, in-vitro studies on the skate ovary have demonstrated that the corpus luteum is also capable of T synthesis and may also contribute to circulating levels of this hormone (Callard et al., 1993). In contrast to E2 and T, the primary source of circulating P4 in female elasmobranchs is corpora luteal tissue, which forms primarily from granulosa cells just prior to or after ovulation (Figure 11.5). The presence of steroidogenic enzymes involved in P4 synthesis has been detected in luteal tissue of S. acanthias and substantial quantities of P4 are produced by luteal minces from both the shark and skate ovary in vitro (Tsang & Callard, 1987a; Fileti & Callard, 1988; Callard et al., 1992). As observed in these studies, production of P4 by the corpus luteum increases with its development, but declines with its age (Tsang & Callard, 1987a; Fileti & Callard, 1988). Granulosa cells from mid- to large-sized ovarian follicles of the skate L. erinacea also are capable of synthesizing P4, but at lower levels than E2 or T, and to a far lesser degree than luteal tissue (Lutton et al., 2005).

Very few studies have examined steroid-binding activity and/or the occurrence of steroid receptors in female elasmobranchs. None-the-less, limited data on this topic suggest that both estrogen receptors (ERs) and progesterone receptors (PRs) are present in most of the major organs that contribute to reproduction, such as the liver and various components of the reproductive tract (Reese & Callard, 1991; Paolucci & Callard, 1998; Koob & Callard, 1999). To the best of the authors’ knowledge, no published studies to date have explored the distribution of androgen receptors (ARs) in female sharks and rays. A more detailed description of putative target organs for gonadal steroids in female elasmobranchs is provided in the following section, along with possible roles for these hormones in reproduction.

3.3. Gonadal Steroid Cycling and Functions 3.3.1. 17b-estradiol (E2) Circulating E2 concentrations generally peak during the period of follicular development in both oviparous and viviparous elasmobranchs (Figure 11.6), suggesting a role for this hormone in regulating synthesis of the yolk protein precursor, vitellogenin (Vtg). The regulation of Vtg production by E2 is well-conserved across nonmammalian vertebrate taxa, and generally occurs in the liver via interactions of E2 with hepatic ERs (Polzonetti-Magni, Mosconi, Soverchia, Kikuyama, & Carnevali, 2004). Following its synthesis, Vtg is transported via the general

Squalus acanthias follicles corpora lutea E ’

ing

P

tch

‘ha

pregnancy

ovulation

parturition 2 years

Sphyrna tiburo E

P

Dasyatis sabina

follicular phase

ovulation 1 year

ro a ibu S. t . sabin D on tati ta cen nema a l p ho trop

pregnancy

parturition

FIGURE 11.6 Patterns of circulating 17b-estradiol (E2) and progesterone (P4) levels during the reproductive cycles of females of three different viviparous elasmobranch species. In Squalus acanthias, P4 levels are high during the first half of pregnancy and then decline to low levels for the second half of pregnancy, when E2 levels are high. In both Sphyrna tiburo and Dasyatis sabina, P4 levels rise during the periovulatory period and remain elevated during early pregnancy, but then decline and remain relatively low for the remainder of pregnancy. 17bestradiol levels rise during late pregnancy in all three species, and, in the placental S. tiburo and aplacental D. sabina, this correlates with a shift in embryonic nutrient supply, indicating that E2 may play an important role in regulating nutrients to developing young. Modified from Koob and Callard (1999), with permission.

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circulation to the ovary, where it is taken up by growing oocytes through the process of receptor-mediated endocytosis. A role for E2 in regulating hepatic Vtg production in sharks and their relatives is supported by evidence for hepatic ERs in the skate L. erinacea (Koob & Callard, 1999), and experimental studies that have demonstrated induction of Vtg production in elasmobranchs in response to E2 treatment (Figure 11.7) (Craik, 1978; Callard, Etheridge, Giannoukos, Lamb, & Perez, 1991; Perez & Callard, 1992; 1993; Prisco et al., 2008b). In addition, increased levels of circulating Vtg coincide with the preovulatory rise in E2 observed in both L. erinacea and S. canicula (Craik, 1978; Craik, 1979; Perez & Callard, 1993). A secondary rise in circulating E2 concentrations also has been observed in the continuous breeder S. acanthias during the latter half of gestation, and appears to regulate hepatic Vtg production and follicular development for the succeeding reproductive cycle. Although recent studies have suggested that the ovarian follicle also may contribute to the production of Vtg in some elasmobranchs (Prisco et al., 2002b), the extent to which this occurs in this group and the importance of E2 in regulating ovarian vitellogenesis is not known. Since the rise in circulating E2 concentrations that occurs during the follicular stage in most female elasmobranchs coincides with increased growth and activity of the oviducal gland, it is possible that E2 also regulates certain aspects of the development and/or function of this organ (Koob, Tsang, & Callard, 1986; Heupel, Whittier, & Bennett, 1999; Gelsleichter, 2004; Sulikowski, Tsang, & Howell, 2004; 2005). This argument is supported by the

FIGURE 11.7 The effects of 17b-estradiol (E2) and progesterone (P4) on plasma vitellogenin in male skates Leucoraja (Raja) erinacea. Intact mature male skates were treated with vehicle, E2, P4, or a combination of E2 and P4, and plasma was assayed for vitellogenin on days 1, 3, 5, and 8. Vitellogenin was detectable in E2-treated animals by day five and rose sharply by day eight. Progesterone treatment in combination with E2 attenuated the E2 response by day eight. Vitellogenin was not detected in control (vehicle) or P4-treated skates. ND, nondetectable. Asterisk indicates p < 0.01 for E2 vs. E2 þ P4 on day eight. Modified from Perez and Callard (1993), with permission.


presence of ERs in the oviducal gland of some elasmobranchs (Reese & Callard, 1991), as well as observations of increased growth and capsule protein synthesis in the oviducal gland of E2-treated female S. canicula (Dodd & Goddard, 1961). However, since no published studies have explored the distribution of ERs among the diverse cell types that make up the oviducal gland, the specific role of E2 in this component of the female reproductive tract remains largely speculative. As for egg fertilization and encapsulation, the passage of fertilized ova from the oviducal gland to the uterus via the isthmus also overlaps with elevations in circulating E2 concentrations that occur in females of some elasmobranch species just prior to and around the time of ovulation. Since ER expression appears to be high in the isthmus (Callard et al., 2005), it is likely that E2 alters its structure in a manner that would favorably influence this transport process. This is well supported by experimental studies, which have demonstrated that E2 can increase the compliance of the isthmus in female S. acanthias (Koob, Laffan, Elger, & Callard, 1983), and the circumference of this compartment in female L. erinacea (Callard & Koob, 1993). Since the isthmus also appears to possess secretory activity, it is possible that E2 regulates other aspects of its physiology that contribute to reproduction. Postovulatory rises in serum E2 concentrations also have been reported to occur during mid to late pregnancy in females of some seasonally breeding elasmobranch species, such as the bonnethead shark and the Atlantic stingray (Figure 11.6) (Manire, Rasmussen, Hess, & Hueter, 1995; Snelson, Rasmussen, Johnson, & Hess, 1997; Tricas, Maruska, & Rasmussen, 2000). Since follicular development for the subsequent breeding cycle does not begin until after parturition in these species, increased levels of E2 may reflect a possible role for this hormone in gestation or parturition. In both cases, increased E2 concentrations coincide with morphological and functional changes in the uterus associated with shifts in embryonic nutrient supply. In the placental viviparous species Sphyrna tiburo, the postovulatory rise in E2 concentrations occurs during formation of placental connections between the gravid female and developing offspring. Similarly, in the aplacental viviparous species D. sabina, E2 levels increase dramatically during a period when the gravid female produces a nutrient-rich uterine histotroph (‘uterine milk’), which nourishes embryos from the middle to late stages of pregnancy. Perhaps E2 plays direct roles in modifying uterine morphology and functions to permit placentation and uterolactation to occur in these species; however, experimental support for this hypothesis is lacking. Lastly, E2 is believed to play an accessory role in modulating uterine contractility and cervical compliance in female elasmobranchs in a manner that would initially retain egg capsules or embryos in the reproductive tract during

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sclerotization or gestation, and later facilitate oviposition or parturition at the appropriate time. This appears to be related to E2’s ability to potentiate the effects of the ovarian-derived peptide hormone relaxin on the uterus and cervix and is discussed further below (see Section 5).

3.3.2. Progesterone (P4) In most female elasmobranchs studied to date, circulating P4 levels increase at or around the time of ovulation and may play a role in suppressing hepatic production of Vtg (Figure 11.6) (Tsang & Callard, 1987b; Fasano, D’Antonio, Pierantoni, & Chieffi, 1992; Manire et al., 1995; Tricas et al., 2000; Mull, Lowe, & Young, 2010). This is supported by the presence of PRs in the liver of female L. erinacea, as well as the ability of P4 to block E2-stimulated vitellogenesis and overall follicular development in this species (see Figure 11.7) (Perez & Callard, 1992; 1993; Paolucci & Callard, 1998). Additional support for this hypothesis comes from studies on S. acanthias, in which E2 treatment was incapable of inducing Vtg production in gravid females during the first half of pregnancy, when endogenous P4 levels remain elevated. This suggests that P4 may restrict Vtg production in female S. acanthias until the second half of pregnancy, when P4 levels decline and E2 levels increase and stimulate yolk production for the subsequent breeding period. More recently, support for P4-regulated inhibition of vitellogenesis also has been observed in the ray Torpedo marmorata, in which circulating P4 concentrations and Vtg expression are inversely correlated (Prisco et al., 2008b). A potential role for P4 in regulating some aspects of the egg-laying process in oviparous elasmobranchs also has been proposed due to the changes in circulating P4 concentrations that occur between ovulation and oviposition in a number of species (Koob et al., 1986; Heupel et al., 1999; Rasmussen, Hess, & Luer, 1999; Sulikowski et al., 2004; Awruch, Pankhurst, Frusher, & Stevens, 2008). This hypothesis is best supported in skates, based on localization of PRs in the reproductive tract of L. erinacea and the observation that P4 treatment can cause early oviposition in this species (Koob & Callard, 1985). Progesterone levels decline in L. erinacea following ovulation and remain low until oviposition occurs, suggesting that P4 may play a role in regulating egg retention (Koob et al., 1986). This is supported by recent studies on the skate Malacoraja senta, in which females without egg cases had higher P4 concentrations than those carrying egg cases of any developmental stage (Kneebone, Ferguson, Sulikowski, & Tsang, 2007). However, in Raja eglanteria, a significant rise in serum P4 concentrations occurs at the time of oviposition, perhaps suggesting an opposing role for P4 in this species (Rasmussen et al., 1999). Based on these findings, as well as the apparent lack of a relationship between endogenous P4 concentrations and egg laying or

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retention in other skate species (i.e., Amblyraja radiata), it is still premature to make generalities about the putative functions that P4 may have in this process.

3.3.3. Androgens As previously mentioned, no published studies to date have investigated the presence of ARs in the reproductive tract of female elasmobranchs. None-the-less, a number of hypotheses regarding the possible functions of androgens in female chondrichthyans have been proposed, based on alterations in circulating T, 11-ketotestosterone (11-KT), or 5a-dihydrotestosterone (DHT) concentrations that have been observed in relation to the reproductive cycle in both oviparous and viviparous species. For example, a preovulatory rise in endogenous T levels occurs in females of several elasmobranch species and likely serves as a precursor for E2 synthesis during this period (Koob et al., 1986; Manire et al., 1995; Snelson et al., 1997; Rasmussen et al., 1999; Tricas et al., 2000). In addition, since increased levels of T coincide with the mating period in some sharks and rays, androgens also have been suggested to play a role in modulating copulatory behavior or aggression in these species (Manire et al., 1995; Rasmussen et al., 1999; Tricas et al., 2000; Mull et al., 2010). In female S. tiburo, the rise in androgen levels that occurs during the mating period persists during the six-month sperm storage period, suggesting that it may regulate certain aspects of this process (Manire et al., 1995). Lastly, based on the increase in serum androgen concentrations in the oviparous skate R. eglanteria, draughtboard shark Cephaloscyllium laticeps, and ratfish Hydrolagus colliei during the egg-laying period, a role for androgens in oviposition and/or encapsulation also has been proposed (Rasmussen et al., 1999; Awruch et al., 2008b; Barnett, Earley, Ebert, & Cailliet, 2009). Although unconfirmed, all of these hypotheses are intriguing and warrant further attention.

4. HORMONAL REGULATION IN MALES 4.1. Testicular Structure and Spermatogenesis The general anatomy of the chondrichthyan male reproductive system has been reviewed in detail by others (Pratt, 1988; Carrier et al., 2004; Engel & Callard, 2005; Jones et al., 2005), and only that information needed to understand the hormonal regulation discussed below will be briefly presented here. The elasmobranch testes are paired structures located anterior to the coelom and suspended from the dorsal body wall by mesorchia. The morphology and functional structure of the testes varies tremendously between species, but can be generally categorized into three different types according to the spatial organization of

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spermatogenesis: (1) diametric or linear testes, found in squalomorph, galeomorph, sphyrnid, and carcharhinid sharks; (2) radial testes, found in lamnoid sharks; and (3) compound testes, found in batoids (Figure 11.4) (Pratt, 1988). In addition to testes, the male reproductive system consists of the associated paired epigonal organs, paired genital ducts (each consisting of efferent ducts, epididymis, and the sperm storage organdthe ampulla epididymis or ductus deferens), the urogenital papilla, the siphon sacs (sharks), the alkaline and clasper glands (batoids), and the intromittent organs, the claspers. Each testis contains two main cell types that differ in embryonic origin and function: germ cells and somatic cells (Leydig and Sertoli cells). The existence of true Leydig cells in chondrichthyan fishes remains controversial (see Section 4.2), but, in those species where they are described, they primarily function in steroid production. Chondrichthyan Sertoli cells are closely associated with the germ cells, and, similar to mammals, likely function to physically support germinal elements, control the microenvironment through secreted products, and serve both as a target and a source of molecules involved in regulating spermatogenesis (Engel & Callard, 2005). However, elasmobranch Sertoli cells differ from those of mammals because they (1) undergo cycles of proliferation, differentiation, and degeneration not seen in mature mammals; (2) are developmentally synchronized with germ cell clones at the same spermatogenetic stage, rather than multiple stages as in mammals; and (3) are primary steroidogenic cells in the testis (Engel & Callard, 2005). Spermatogenesis in chondrichthyan fishes occurs within a functional testicular unit called a ‘spermatocyst’ (or sometimes described as a ‘follicle,’ ‘ampulla,’ or ‘lobule’). The spermatocyst is an anatomically discrete sphere bounded by an acellular basal lamina that consists of a developing syncytial germ cell clone and its associated Sertoli cells. Spermatocysts are formed when a single spermatogonium in the germinal zone of the testis associates with a pre-Sertoli cell. The germ celleSertoli cell spermatocyst unit then undergoes a sequence of complex cellular events (i.e., mitosis, meiosis, apoptosis, maturation) that is very similar to the spermatogenetic process in all vertebrates, which ultimately results in formation of mature sperm that are released into the efferent ductules when the spermatocyst bursts.

4.2. Steroidogenesis, Steroidogenic Enzymes, and Steroid Receptors The exact location of testicular steroid synthesis in chondrichthyans is still debated. In most male vertebrates, gonadal steroids are produced by cells that lie outside of the germ cell or testicular spermatocyst compartments (i.e., the


interstitial or Leydig cells). Early studies described Leydiglike cells in the interstitial tissue of the dogfish shark, but they were portrayed as undifferentiated and did not undergo structural changes associated with spermatogenesis (Pudney & Callard, 1984b). More recent studies have demonstrated that true Leydig cells do exist in elasmobranchs, but that Sertoli cells are likely the more ancestral producers of steroid hormones (Pudney & Callard, 1984b; Callard, Mak, DuBois, & Cuevas, 1989; Prisco et al., 2008a). Elasmobranch Sertoli cells contain both cytological and enzymatic characteristics of steroid-producing cells and are thought to be the major source of androgens in these fishes (Simpson & Wardle, 1967; Pudney & Callard, 1984a; Dubois & Callard, 1989; Engel & Callard, 2005). This is supported by studies in the dogfish that show increased activity of key enzymes that control androgen biosynthesis in Sertoli cells as spermatogenesis proceeds (Callard, Pudney, Mak, & Canick, 1985). However, in both the dogfish (McClusky, 2005) and the ray T. marmorata (Prisco et al., 2002a; 2003; 2008a), androgens are now believed to be produced by several different cell types: Leydig cells, Sertoli cells, and possibly germ cells (i.e., spermatogonia), and the relative role(s) of Leydig and Sertoli cells in steroid production appears to change during spermatogenesis. Immunocytochemical examination of the Torpedo ray testis has shown that key enzymes involved in the T biosynthetic pathway, such as 3b-hydroxysteroid dehydrogenase (3bHSD) and 17b-hydroxysteroid dehydrogenase (17b-HSD), are expressed in Sertoli cells before meiosis and after spermiation, and in Leydig cells only during meiosis (Prisco et al., 2008a). Activity of the enzyme aromatase (P450aro ¼ CYP19) involved in converting T to E2 is higher in elasmobranch testis regions that are undergoing meiosis compared with both less mature and more mature areas (Callard et al., 1985). Although P450aro has been cloned in several elasmobranch species (Ijiri, Berard, & Trant, 2000; Engel & Callard, 2005), its role during spermatogenesis and other aspects of male reproduction remains unknown. In contrast to P450aro, the enzyme that converts T to DHT (i.e., 5a-reductase) is higher in regions with premeiotic spermatocysts compared with both postmeiotic and meiotic stages (Cuevas, Collins, & Callard, 1993), which is also consistent with the presence of DHT in premeiotic areas following in-situ perfusion of the dogfish testis with [3H]-T (Callard et al., 1989; Cuevas & Callard, 1992). There are relatively few published studies on testicular steroid receptors in chondrichthyan fishes, and virtually all have been performed in the dogfish Squalus (Callard & Mak, 1985; Callard et al., 1985; Cuevas & Callard, 1992). Androgen-binding activity in Squalus testis was highest in regions with premeiotic-stage spermatocysts (Cuevas & Callard, 1992), and immunocytochemistry showed ARs localized to somatic cells (i.e., Sertoli cells) but not germ

Chapter | 11


cells (Engel & Callard, 2005). This stage-related pattern of AR expression in premeiotic regions of the dogfish testis was later confirmed by measurement of AR mRNA levels (Engel & Callard, 2007), and implies that androgens serve an important function during early spermatogenesis. Androgen receptor has also been recently cloned from the testis of the brown-banded bambooshark Chiloscyllium punctatum, and, similar to other vertebrates, classical androgens (T, 11-KT, DHT) binding to this shark AR have been shown to activate target genes via an androgen response element (Ogino, Katoh, Kuraku, & Yamada, 2009). Estrogen receptors in the dogfish testis are also highest in regions with stem cells, spermatogonia, and premeiotic stages, but virtually absent in areas of mature germ cells (Callard et al., 1985; Ruh, Singh, Mak, & Callard, 1986; Engel & Callard, 2005; 2007). In contrast, P450aro was localized to more mature meiotic regions, and, because the blood flow in the dogfish testis is directed from more advanced to less advanced spermatocyst stages (Cuevas, Miller, & Callard, 1992), E2 may play a paracrine role to signal the advance of spermatogenetic development (Engel & Callard, 2007). The prevalence of both AR and ER in early premeiotic stages indicates that androgens and estrogens may cooperate to regulate gene expression prior to meiosis in the shark testis (Callard, 1991). Progesterone receptors also have been described in dogfish testes (Cuevas & Callard, 1992), but, in contrast to AR and ER, their distribution is associated with regions of mature or late-stage spermatocysts and thus P4 may function in spermiogenesis and sperm release. Progesterone receptor mRNA expression later confirmed these stagedependent binding studies, but also revealed high expression of PRs in premeiotic stages (Engel & Callard, 2007). In addition to the steroid receptors mentioned above, a cytosolic nonreceptor-binding protein has been characterized from subfractions of dogfish testis (Mak & Callard, 1987). Based on its high affinity, broad specificity, molecular weight, isoelectric point, and dimeric structure, it was thought to be related to androgen-binding protein (ABP), found in mammalian testis. The concentration of this putative ABP increases as spermatogenesis proceeds, which is coincident with Sertoli cell development and androgen biosynthesis, and thus ABP may serve as a testicular steroid reservoir in sharks.

4.3. Gonadal Steroid Cycling and Functions Numerous gonadal steroids have been described in male elasmobranch fishes (Callard, 1988; Garnier, Sourdaine, & Jegou, 1999; Manire, Rasmussen, & Gross, 1999), but only a few have been investigated in detail in relation to the reproductive cycle (T, DHT, 11-KT, E2, and P4) and will be discussed below. Further, little is known about circulating gonadal steroid levels in male holocephalans (but see

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Barnett et al., 2009) and therefore all of the information discussed below refers primarily to elasmobranchs.

4.3.1. Androgens Testis development and seasonal spermatogenesis both appear to be regulated by androgens. Serum T levels (and/ or other androgens such as DHT) are often elevated during the middle to late stages of spermatogenesis, which is coincident with the presence of mature spermatocysts in the testes (Manire & Rasmussen, 1997; Snelson et al., 1997; Heupel et al., 1999; Tricas et al., 2000; Sulikowski et al., 2004; Awruch et al., 2008b). Serum T levels also increase with sexual maturation and clasper length, and are elevated during the mating season in some elasmobranch species. Therefore, androgens may have effects on the development of both secondary sex characteristics and copulatory and aggressive behaviors, as seen in other vertebrates. Serum androgen concentrations also are elevated during periods of increased semen transport in sharks and batoids (Manire & Rasmussen, 1997; Garnier et al., 1999; Heupel et al., 1999; Tricas et al., 2000), and thus may influence development and function of the gonoducts and/or maturation of sperm. The seminal fluid of some sharks also contains high concentrations of steroids and steroidogenic enzymes (Simpson, Wright, & Gottfried, 1963; Simpson, Wright, & Hunt, 1964) that may play an important yet undescribed role in regulation of the male reproductive tract, semen transport and storage, and/or copulation and fertilization. In seasonally breeding elasmobranchs, T is often correlated with gonadal recrudescence, final sperm maturation, and the onset of copulatory activity. However, studies that have examined both circulating steroids and histological stage of the testes indicate that circulating T levels more closely reflect spermatogenetic stage than overall testis mass or gonadosomatic index (GSI) (Figure 11.8) (Heupel et al., 1999; Tricas et al., 2000; Sulikowski et al., 2004; Mull, Lowe, & Young, 2008). This is also confirmed by the presence of stage-specific steroidogenic enzymes in the testis, and high levels of T produced by cultures of more mature spermatogenetic stages (Sourdaine et al., 1990; Sourdaine & Garnier, 1993; Engel & Callard, 2005). In species that are capable of reproducing year-round, however, T levels remain relatively constant throughout the year and there is only a weak (or absent) correlation between circulating T concentration and the number of late-stage spermatocysts (Kneebone et al., 2007). Future studies that account for the different reproductive modes and breeding cycles found in chondrichthyans are needed to fully interpret the relationship between T, spermatogenesis, and male reproductive competence. 11-ketotestosterone is thought to be the main androgen in teleost fishes, but its function in male elasmobranchs is

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FIGURE 11.8 Stages of testes development and changes in circulating steroid levels in male Atlantic stingrays (Dasyatis sabina). Left side shows four histological stages of spermatogenesis expressed as a percentage of total gonad weight for each monthly sample (closed circles), plotted with steroid concentrations for testosterone (T) and 17b-estradiol (E2) expressed in relative values. Right side shows representative histological sections of the testis from each stage. The first peak in circulating T levels is coincident with spermatocyte presence, while the second peak is coincident with a relatively brief activity of spermatocytes and the peak of sperm maturation. Peak circulating E2 levels are associated only with spermatocyte production. IS, immature sperm; MS, mature sperm; SC, spermatocytes; SE, Sertoli cells; ST, spermatids. Scale bars ¼ 50 mm. (left panel) Modified from Tricas et al. (2000). (right panel) Modified from Maruska, Cowie, and Tricas (1996), with permission.

less clear. Circulating 11-KT has been detected in the viviparous bonnethead shark S. tiburo (Manire et al., 1999) and stingray U. halleri (Mull et al., 2008), and the oviparous dogfish S. canicula (Garnier et al., 1999), where it may contribute to testicular development. However, 11-KT was undetectable in another oviparous shark, C. laticeps (Awruch et al., 2008b). In studies where both T and 11-KT were examined, serum T levels were higher but the patterns of both androgens were very similar. Serum levels of the androgen DHT also parallel those of T in most species, and DHT is thought to be an important reproductive hormone in male elasmobranchs. Plasma levels of T, DHT, and 11-KT within a single species are only known for the placental viviparous bonnethead (Manire et al., 1999) and oviparous dogfish sharks (Garnier et al., 1999), and future studies are needed to determine the relative roles of these different androgens in male chondrichthyan reproduction.

4.3.2. Estrogens The role of E2 in male elasmobranch reproduction is still enigmatic, partially because it is often not measured in males. Many elasmobranch species show distinct seasonal patterns in circulating E2 concentrations that are related to early and middle stages of spermatogenesis (Manire & Rasmussen, 1997; Snelson et al., 1997; Tricas et al., 2000; Sulikowski et al., 2004). However, other studies show either little or irregular seasonal variation in circulating E2 levels in males (Garnier et al., 1999; Awruch et al., 2008b), which makes any generalizations on function premature until more species with different testicular organization and breeding cycles are examined. In male Atlantic stingrays, plasma E2 levels follow testis growth and primary androgen increases during peak spermatocyte activity, but E2 levels are not elevated during a secondary androgen increase that

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occurs in this species (see Figure 11.8) (Tricas et al., 2000). Aromatase enzyme activity has been shown to be highest in primary and secondary spermatocytes of the dogfish (Callard et al., 1985), and thus may contribute to the elevated E2 levels during times of high circulating T in the stingray and other male elasmobranchs. Estrogen receptors are also localized to testicular regions that contain premeiotic spermatocysts (Callard et al., 1985; Callard, 1992), and E2 may have paracrine effects on germ cells in earlier stages of spermatogenesis. Treatment of these premeiotic germ cells with E2 results in a dose-dependent reduction in both cell proliferation and programmed cell death, which indicates it may regulate spermatogenesis via a negative feedback system on developmental arrest (Betka & Callard, 1998). Elevated E2 concentrations also coincide with increased cell proliferation and growth in the epididymis and seminal vesicle of male Atlantic stingrays (Gelsleichter, 2004), which indicates it may play a role in genital tract function similar to that described for mammals (Hess, 2003; Shayu, Hardy, & Rao, 2007).

4.3.3. Progesterone (P4) In male elasmobranchs, P4 is thought to be a substrate for androgen synthesis during later stages of spermatogenesis, or it may play a role in the regulation of spermiogenesis and/or spermiation (Gelsleichter, 2004). The androgen precursor role for P4 stems from the fact that circulating P4 levels coincide with testicular development and mirror the concentrations of T and DHT in some male elasmobranchs (Rasmussen & Gruber, 1990; 1993; Manire & Rasmussen, 1997; Tricas et al., 2000). However, other studies show no correlation between P4 and androgen concentrations (Snelson et al., 1997; Garnier et al., 1999; Awruch et al., 2008b). The increase in P4 levels during later stages of spermatogenesis in some species (Gelsleichter et al., 2002), and the observation that PRs show greater expression in late-stage (postmeiotic) compared with early-stage spermatocysts (Cuevas & Callard, 1992), support the hypothesized role of P4 in spermiogenesis and/or spermiation. In addition, the higher P4 concentrations in males compared with females of some elasmobranch species (Manire & Rasmussen, 1997; Rasmussen et al., 1999) indicate some important role in male reproductive physiology that warrants future study.

5. OTHER HORMONES INVOLVED IN REPRODUCTION IN MALES AND FEMALES 5.1. Corticosterone (CORT) Serum concentrations of the steroid hormone corticosterone (CORT) show sex-specific and seasonal variations associated with reproduction in several elasmobranchs

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(Snelson et al., 1997; Manire, Rasmussen, Maruska, & Tricas, 2007). In bonnethead sharks, serum CORT levels covary with gonadal steroids in both sexes; are correlated with testicular growth, spermatogenesis, and mating in males; and are correlated with vitellogenesis, sperm storage, migration, mating, and early pregnancy in females. In the stingray D. sabina, male CORT levels were similarly elevated during testis development and mating, but, in contrast to the bonnethead, female CORT levels were correlated with late pregnancy, parturition, and postpartum stages (Figure 11.9) (Manire et al., 2007). These studies indicate that CORT plays some important role in seasonally breeding elasmobranchs possibly related to the stress axis, but its function may differ between the sexes and among species with different reproductive modes.

5.2. Relaxin The peptide hormone relaxin is in the insulin superfamily, and is best known for its role in preparing the female reproductive tract for parturition. Relaxin has been detected in the ovaries of both sharks and batoids (Gowan et al., 1981; Reinig et al., 1981; Bullesbach et al., 1986; Bullesbach, Schwabe, & Callard, 1987; Steinetz, Schwabe, Callard, & Goldsmith, 1998), and may play a role during pupping (birth) and/or egg-laying in elasmobranchs. For example, relaxin can increase cervical cross-sectional area in late-stage pregnant S. acanthias (Koob, Laffan, & Callard, 1984), and it increases compliance of the female reproductive tract including the cervix in the skate L. erinacea (Callard et al., 1993). These effects are similar to the ability of relaxin to increase the circumference of the mammalian birth canal (Steinetz, O’Byrne, Butler, & Hickman, 1983). Relaxin also inhibits uterine contractions in the dogfish, which would protect the encapsulated embryos during early pregnancy and prevent early parturition (Figure 11.10) (Sorbera & Callard, 1995), and, as previously mentioned, these effects are potentiated by E2. Relaxin is also produced by male reproductive structures in several vertebrates; is found in the testis, blood, and semen of some elasmobranchs (Steinetz et al., 1998; Gelsleichter, Steinetz, Manire, & Ange, 2003); and is hypothesized to play a role in regulating male fertility (Weiss, 1989). Serum relaxin concentrations are elevated specifically during late spermatogenesis and the copulatory period in male S. tiburo (Figure 11.10) (Gelsleichter et al., 2003), and relaxin levels in the semen are 1000 times higher than in the circulation (Gelsleichter, 2004). These data, along with the presence of a relaxin-like compound (raylaxin) produced by the alkaline gland of skates and rays (Bullesbach, Schwabe, & Lacy, 1997), indicate relaxinrelated compounds may regulate semen quality or sperm motility, or perhaps facilitate insemination by controlling uterine contractility in postmated females.

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Volkoff et al., 1999). Elevated circulating thyroid hormones during the period of ovulation, gestation, and maternalefetal nourishment in both the stingray D. sabina and bonnethead shark suggest increased production of these hormones may relate to greater metabolic demand during the energetically costly processes used to provide nutrients to developing embryos (Volkoff, Wourms, Amesbury, & Snelson, 1999; McComb, Gelsleichter, Manire, Brinn, & Brown, 2005). Maternal serum and yolk levels of T3 and T4 in the bonnethead shark increased from the preovulatory to postovulatory periods, and peaked during pregnancy (Figure 11.11) (McComb et al., 2005). Further, higher yolk TH levels were found in a bonnethead population in Tampa Bay that shows faster rates of embryonic development compared with a Florida Bay population with slower development, indicating that thyroid hormones may play some role in controlling the rate of embryonic development in these animals (Figure 11.11) (McComb et al., 2005).

5.4. Calcitonin (CT)

FIGURE 11.9 Monthly plasma corticosterone (CORT) concentrations in male and female Atlantic stingrays (Dasyatis sabina) in relation to reproductive events. Corticosterone levels in females were elevated during late pregnancy, parturition, and postpartum stages, while male CORT levels were elevated during spermatogenesis and the protracted mating season. Bars show medians and 25the75th quartiles (error bars), and sample sizes are indicated above each bar. Lines above bars indicate groups that were significantly different from remaining groups (KW ANOVA, p < 0.05). Modified from Manire et al. (2007), with permission.

5.3. Thyroid Hormones The thyroid hormones triiodothyronine (T3) and thyroxine (T4) are thought to interact with the HPG axis to regulate vertebrate reproduction, and serum levels are correlated with reproductive and developmental events in several elasmobranch species (Crow, Ron, Atkinson, & Rasmussen, 1999; Gash, 2000; McComb et al., 2005;

The polypeptide hormone calcitonin (CT) is thought to regulate certain aspects of reproduction in many vertebrates, including pregnancy, follicular development, and embryonic development (Zaidi, Inzerillo, Moonga, Bevis, & Huang, 2002). Calcitonin is produced by the ultimobranchial gland in elasmobranchs, a paired organ embedded in the musculature between the pharynx and pericardial cavity. In the stingray Dasyatis akajei, ERs were localized to the ultimobranchial organ and E2 increased CT production (Takagi, Suzuki, Sasayama, & Kambegawa, 1995; Yamamoto, Suzuki, Takahashi, Sasayama, & Kikuyama, 1996). In female bonnethead sharks, serum CT concentrations showed a temporal pattern where peak levels occurred during the yolk-dependent stage of pregnancy, and CT-ir was localized to the duodenum and pancreas of developing embryos during this same stage, suggesting it may be involved in digestion of yolk and fetal nutrition in this placental species (Figure 11.12) (Nichols, Gelsleichter, Manire, & Cailliet, 2003).

5.5. Serotonin (5-HT) High concentrations of serotonin (5-HT) have been found in siphon sac secretions of mature male spiny dogfish S. acanthias, but 5-HT was either absent or found only in trace amounts in the semen of immature males (Mann, 1960). Siphon sacs in mature males had 200 times more 5-HT than immature males, and siphon sac secretions stimulated uterine contractions in vitro in S. acanthias (Mann, 1960; Mann & Prosser, 1963). These data indicate that, during copulation, 5-HT from the male may cause

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FIGURE 11.10 Relaxin function in female and male elasmobranch reproduction. (a) Effect of homologous Squalus relaxin (sRlx) on spontaneous myometrial contractions and intrauterine pressure cycles in the female dogfish Squalus acanthias. In-vitro myometrial contraction recordings from a uterine tissue strip of a stage C dogfish (ovary contains large 30e34 mm diameter follicles and 12e20 cm embryos free in uterine lumen) before (control) and after intravenous injection of sRlx. Squalus relaxin significantly decreased the frequency of contractions in a dose-dependent manner by eliminating smaller low-intensity contractile activity interspersed between the larger contractions. Contraction patterns then returned to control patterns after washout. Intrauterine pressure cycles recorded from a stage C dogfish in vivo showed that Rlx caused a dose-dependent decrease in mean frequency of contractions. Thus, sRlx slows the frequency of uterine contractions in third-trimester sharks, a time when progesterone (P4) is reduced and 17bestradiol (E2) levels are rising. (b) Mean serum relaxin concentrations in relation to gonad size (testes width multiplied by length) and reproductive events in mature male bonnethead sharks (Sphyrna tiburo). Serum Rlx concentrations were significantly increased during late spermatogenesis and mating (i.e., September and October levels were greater than all other months except November). (a) Reproduced from Sorbera and Callard (1995). (b) Reproduced from Gelsleichter et al. (2003), with permission.

uterine contractions in the female that aid in sperm transport and fertilization (Mann & Prosser, 1963). Serotonin is also a ubiquitous neurotransmitter in the brain that plays a role in sexual behaviors and aggression in many vertebrates. The embryonic and ontogenetic development of the primary 5-HT system in the brain of S. canicula was recently described as similar to mammals (Carrera, Molist, Anadon, & Rodriguez-Moldes, 2008), suggesting that conserved functions of 5-HT in reproductive-related behaviors may extend to chondrichthyans as well.

5.6. Neurohypophysial Hormones Neurohypophysial hormones are a family of structurally and functionally related nonapeptides that include the vasopressin and oxytocin homologs. Chondrichthyans contain the arginine vasopressin homolog, arginine vasotocin (AVT), as well as at least eight types of oxytocinfamily peptides identified in different species (i.e., oxytocin, isotocin, glumitocin, valitocin, aspargtocin, asvatocin, phasitocin, phasvatocin) (Gwee, Tay, Brenner, & Venkatesh, 2009). Arginine vasotocin is an important

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(c)

10 Florida Bay

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Yolk T4 concentration (ug/dl)

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PrePostPregnancy Ovulatory Ovulatory

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(d) Florida Bay Tampa Bay

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15 10

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(b) Yolk T3 concentration (ng/ml)

*

0.1

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5

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0.0

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Reproductive stage

FIGURE 11.11 Yolk (a, b) and maternal serum (c, d) thyroid hormone concentrations during different reproductive stages of female bonnethead sharks (Sphyrna tiburo) sampled from two different sites (Florida Bay and Tampa Bay, FL). Asterisks indicate significant differences between reproductive stages within a site, and illustrate increases in thyroid hormone levels from preovulatory to pregnancy stages. Triangles indicate significant differences between sites during a specific reproductive stage, which may be related to reported differences in embryonic development rates between these sites. T3, triiodothyronine; T4, thyroxine. Reproduced from McComb et al. (2005), with permission.

regulator of reproductive and social behaviors across vertebrate taxa, and its distribution in the central nervous system is relatively well conserved (Goodson & Bass, 2001). The distribution of AVT-ir neurons and fibers in the brain is known for only a single chondrichthyan fish, the dogfish S. canicula (Vallarino, Viglietti-Panzica, & Panzica, 1990). While nothing is known of the reproductive physiological function of AVT in chondrichthyans, the peptide distribution in the dogfish brain is similar to that of other fishes and tetrapods and consistent with roles as both a hypophysiotropic molecule influencing the hypothalamicepituitaryeadrenal (HPA) axis and a neuromodulator of reproductive-related functions (Vallarino et al., 1990; Goodson & Bass, 2001). In mammals, oxytocin influences smooth muscle contraction in the ovary and uterus and plays a role in ovulation and parturition. A recent study also detected both AVT and oxytocin in the ovary of the holocephalan elephant shark (Callorhinchus milii) during the peak spawning season, suggesting a paracrine role for these hormones in ovulation and parturition (Gwee et al., 2009).

6. HORMONES, SEXUAL DIFFERENTIATION, AND SEXUAL MATURATION Although studies on sexual differentiation and sex determination in chondrichthyan fishes are limited, the mechanisms involved are different from those of teleosts, but similar to those in amphibians and amniotes (Hayes, 1998). Sex determination is relatively stable in chondrichthyans, as transplantation studies show that gonadal tissue differentiates into ovary or testis according to its genetic sex and is independent of the sex of the transplantation host (Thiebold, 1964). However, as in all vertebrates, development of the reproductive system in chondrichthyan fishes is also influenced by endogenous steroid hormones. Chieffi (1967) observed feminization of embryonic gonads following injections of E2, P4, T, and deoxycorticosterone into the external yolk supply of embryonic Torpedo rays, and similar effects were seen after T and E2 injections in the dogfish S. canicula (Thiebold, 1953; 1954). During pregnancy, E2 may

Chapter | 11


FIGURE 11.12 Maximal ova size (a) and serum calcitonin concentrations (b) in relation to reproductive events in mature female bonnethead sharks (Sphyrna tiburo). Serum calcitonin concentrations during early pregnancy are significantly greater than at all other stages except the postovulatory period (p < 0.05), suggesting an important role in embryonic development. Values displayed are means þ standard error of mean (SEM) for maximal ova size, and medians þ semi-interquartile range for serum calcitonin concentrations. Values in parentheses represent sample sizes for each reproductive stage. Reproduced from Nichols et al. (2003), with permission.

regulate structures that provide nutrients to the developing young, such as hepatic yolk synthesis in oviparous species, and placental or trophonemata growth and function in viviparous species (Callard et al., 2005). Steroid hormones (E2, P4, T) also are transferred from mother to early-stage embryos via yolk provisions in the bonnethead shark, and E2 especially may be involved in sexual differentiation and the initiation of embryonic steroidogenesis (Figure 11.13) (Manire, Rasmussen, Gelsleichter, & Hess, 2004).

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FIGURE 11.13 Yolk (a) and serum (b) 17b-estradiol (E2) concentrations during reproductive stages from preovulatory to preimplantation (preplacental) stages of the female placental viviparous bonnethead shark (Sphyrna tiburo). Yolk E2 concentrations were greater than serum concentrations in postovulatory and early pregnancy stages. Preovulatory E2 serum concentrations were also significantly less than yolk E2 concentrations during ovulatory, postovulatory, and preimplantation stages, and greater than yolk E2 concentrations during early pregnancy. The decline in yolk E2 levels between postovulatory and early pregnancy is consistent with E2 utilization during embryonic development, and the subsequent dramatic increase in the postimplantation stage may be due to hormone production by the embryo; possibly concurrent with or following sex differentiation. Data are plotted as medians with 25th and 75th percentiles (error bars), and sample sizes are indicated above bars. Reproduced from Manire et al. (2004), with permission.

Sexual maturation (or puberty) is associated with activation of the HPG axis in most vertebrates, and chondrichthyan fishes are likely no exception. Elevated levels of circulating steroid hormones appear to be

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essential for development of gonads, reproductive tract, and accessory sex organs (e.g., claspers); mating behavior; and feedback regulation of the brain and pituitary. This is supported by studies that show higher levels of circulating gonadal steroids in mature vs. immature elasmobranchs (Rasmussen & Murru, 1992; Rasmussen & Gruber, 1993; Manire et al., 1999; Gelsleichter et al., 2002). Only a few studies have coupled measurements of circulating gonadal steroid levels with morphological and histological data to more precisely determine the endocrine correlates and age at sexual maturity in sharks (Gelsleichter et al., 2002; Awruch, Frusher, Pankhurst, & Stevens, 2008; Awruch et al., 2008b), skates (Sulikowski et al., 2005; 2006), and holocephalans (Barnett et al., 2009). These studies show concurrent increases in spermatogenesis, clasper length, and plasma T concentrations associated with sexual maturity in males, and increased E2 levels positively correlated with ovary mass, follicle size, and shell gland mass at sexual maturity in females (Figure 11.14).


Several studies in male elasmobranchs show increased serum androgen concentrations as sexual maturation and clasper length increase (Garnier et al., 1999; Heupel et al., 1999; Awruch et al., 2008a; 2008b). However, androgen sensitivity of the claspers and hormonal regulation of these organs are relatively unstudied. A small increment of clasper growth has been observed following androgen injection or implantation in immature skates and dogfish (Hisaw & Abramowitz, 1938; Dodd, 1960). In contrast, there was no direct relationship between androgen concentration and clasper elongation during puberty in S. tiburo (Gelsleichter et al., 2002), and T treatment of clasper cartilage explants from pubertal male bonnethead sharks had no effect on their growth (Gelsleichter, 2004). Hypophysectomy and T treatment also had no effect on in-vivo clasper growth in immature male elasmobranchs (Wourms, 1977). These apparently contradictory findings highlight the need for future studies on additional species, as well as examination of the role of aromatization on clasper growth, since E2 and the growth

FIGURE 11.14 Relationship between sexual maturity, morphology, and steroid hormone levels in male and female winter skates (Leucoraja ocellata). Morphological, histological, and steroid hormone levels were used to assess size and age at sexual maturity: 50% maturity for males was at 730 mm total length and 11 years; 50% maturity for females was at 760 mm total length and 11e12 years. (a) Clasper length and testis mass as males progress through sexual maturity and increase in total length. (b) Proportion of mature spermatocysts and circulating testosterone (T) levels as males progress through sexual maturity. (c) Ovary and shell (nidamental or oviducal) gland mass as females progress through sexual maturity and increase in total length. (d) Follicle diameter and circulating 17b-estradiol (E2) concentrations as females progress through sexual maturity. Average age is given above each representative size class. Values expressed as mean standard error of mean (SEM). Modified from Sulikowski et al. (2005), with permission.

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hormone-insulin-like growth factor I axis is thought to mediate the effect of T on pubertal growth of the mammalian skeletal system (Grumbach, 2000), and thus may operate similarly in chondrichthyan species.

7. HORMONES, REPRODUCTIVE BEHAVIORS, AND SENSORY FUNCTION Reproductive behaviors, including those associated with aggression and territorial defense, courtship and mating, and parental care, are influenced by or correlated with gonadal steroids in many vertebrates. In contrast, there are very few examples of the relationship between reproductive behaviors and gonadal steroids in chondrichthyan fishes. Correlational studies show that circulating androgen levels often are elevated during the mating season in some male elasmobranchs (Heupel et al., 1999; Rasmussen et al., 1999; Tricas et al., 2000), and possibly are associated with aggression and courtship behaviors, but steroid profiles in other species show androgen peaks that occur several months prior to mating (Manire & Rasmussen, 1997; Henningsen, Murru, Rasmussen, Whitaker, & Violetta, 2008). Direct observations of elasmobranch mating events in the wild are rare, and thus there is little information on steroid profiles from actively courting individuals, without which transient increases in circulating steroid levels due to social interactions or the act of copulation would not be detected. However, Rasmussen and Gruber (1990) did find elevated levels of circulating E2 and T in female lemon sharks during active courtship. Henningsen et al. (2008) showed that some components of sexual conflict behaviors such as increased swimming speed, reduced interest in food, tailing (one male follows another male so closely that the lead shark’s tail movement is restricted), and male dominance biting were associated with steroid profiles in male sandtiger sharks (C. taurus), whereas other behaviors such as nosing, following, and copulation attempts were not. Androgen levels within individual sharks also were related to their social position in the dominance hierarchy, with lower levels found in subordinate animals, possibly due to androgen-induced reproductive suppression (Henningsen et al., 2008). In Atlantic stingrays (D. sabina), androgens shift the frequency tuning of the electrosensory system, thus allowing males to better detect the bioelectric fields of buried females during the breeding season (Sisneros & Tricas, 2000). This finding is confirmed by both the natural seasonal cycling of androgens in this species (Tricas et al., 2000) and laboratory implants of the nonaromatizable androgen, DHT (Sisneros & Tricas, 2000). Steroid action on the electrosensory system is further supported by preliminary studies that localized ARs to the peripheral electrosensory ampullae of Lorenzini (Sisneros, 1999).

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As mentioned above, GnRH also may influence the sensitivity of visual, auditory, mechanosensory, electrosensory, and olfactory systems to facilitate mate detection and copulatory behaviors, but this requires further study. Courtship in chondrichthyan fishes often includes a precopulatory behavior called ‘following’ (also termed ‘close follow,’ ‘parallel swimming,’ or ‘chasing’), where the male closely follows the female (within ~one body length) (Carrier et al., 2004; Pratt & Carrier, 2005). This behavior coupled with the existence of a well-developed olfactory system in most elasmobranchs (Schluessel, Bennett, Bleckmann, Blomberg, & Collin, 2008) has led to the hypothesis that pheromones or hormone metabolites are released by the female to trigger or initiate sexual behavior and readiness in males (Johnson & Nelson, 1978; Demski, 1990; Pratt & Carrier, 2001; Hueter, Mann, Maruska, Sisneros, & Demski, 2004). However, there are currently no published experimental studies on pheromones or the use of olfaction during reproductive behaviors in chondrichthyans. Given the importance of olfactory cues during reproduction demonstrated in both higher (e.g., bony fishes, amphibians, reptiles, and mammals) and lower (e.g., lampreys) vertebrate taxa coupled with a legendary olfactory sensing capability, this area of research deserves future attention in chondrichthyans.

8. ENVIRONMENTAL INFLUENCES ON CIRCULATING HORMONE LEVELS AND REPRODUCTION Water temperature and photoperiod are common environmental cues used by seasonally breeding fishes to synchronize reproductive activities and ensure successful mating. Many elasmobranchs are seasonal breeders, and are hypothesized to use seasonal changes in water temperature and/or day length to coordinate reproductive physiology, gametogenesis, and behavior. There is some correlation between circulating steroid hormones and water temperature and/or day length in both males and females of several elasmobranch species (Garnier et al., 1999; Heupel et al., 1999; Rasmussen et al., 1999; Tricas et al., 2000; Mull et al., 2008; 2010). For example, circulating androgen (T and/or 11-KT) concentrations in males were negatively correlated with ambient water temperature and photoperiod (Garnier et al., 1999; Heupel et al., 1999; Mull et al., 2008), and an experimental increase in water temperature from ambient (18e20 C) up to 25 C caused a 15-fold decrease in plasma T levels in male round stingrays (Figure 11.15) (Mull et al., 2008). Steroid production in testicular tissue was also shown to be temperature- but not photoperioddependent in the dogfish S. canicula (Dobson & Dodd, 1977c; Kime & Hews, 1982). Although limited, these studies indicate that temperature and/or day length

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Environmental contaminants (e.g., heavy metals, dioxins, polychlorinated biphenyls, polycyclic aromatic hydrocarbons) can also potentially alter endocrine function and possibly reproductive fitness in some chondrichthyan fishes (see Chapter XX, this volume). In freshwater Atlantic stingrays, serum steroid levels were elevated in subpopulations sampled in a high organochlorine pesticide (OC)-contaminated site compared to intermediate and low OC sites, but there was no evidence of reproductive impairment (Gelsleichter, Walsh, Szabo, & Rasmussen, 2006). The heavy metal cadmium (Cd) is a strong spermatotoxicant that increases germ cell apoptosis, causes spermiation failure, and compromises bloodetestis barrier function in mammals, but also has been shown to have similar effects and accumulate in a stage-specific manner (premeiotic > meiotic > postmeiotic) in the elasmobranch testis (Betka & Callard, 1999; McClusky, 2006; 2008). Although not yet directly tested in elasmobranchs, Cd is a known endocrine-disrupter in other vertebrates, where it mimics estrogen and has sex steroid receptor-activation and -inhibition effects. These detrimental effects on endocrine and reproductive functions are worrisome given the abundance of elasmobranch species that inhabit often heavily polluted estuarine and inshore waters.

9. CONCLUSIONS AND FUTURE DIRECTIONS

FIGURE 11.15 Relationship between temperature and photoperiod and circulating testosterone (T) concentrations in the round stingray (Urobatis halleri). Plasma T levels were negatively correlated with both day length (a) and water temperature (b). (c) Plasma T levels also decreased 15-fold when the water temperature was experimentally raised from 18e20 C to 25 C, and then increased again when it was reduced to 14 C. Bars represent mean þ standard error of mean (SEM) and letters indicate significant groupings (a ¼ 0.05). Black bar shows the control group, C, which was not subject to water temperature manipulations. Modified from Mull et al. (2008), with permission.

probably play important roles in the regulation of circulating steroid levels and reproduction in seasonally breeding chondrichthyan fishes, which is similar to that observed in other vertebrate taxa, but the relative roles of different environmental cues likely depend on species and habitat.

Studies on chondrichthyan reproductive endocrinology have provided important information on how hormones regulate reproduction in this diverse and successful group of fishes, and highlight the fact that many regulatory mechanisms are conserved through vertebrate evolution. However, despite their critical phylogenetic position, there is still a paucity of information on the function of certain hormones in regulating various aspects of chondrichthyan reproduction when compared to most other vertebrate groups. The diversity of breeding strategies and maternalefetal nutritional modes found in sharks, skates, rays, and chimaeras also makes it difficult to generalize about hormonal regulation of reproduction. The majority of information on reproductive hormone function is centered on only a few species (e.g., S. canicula, S. acanthias, L. erinacea, D. sabina, S. tiburo). Although these species do represent different reproductive modes, they are only a limited sampling, and future studies should use the comparative approach to include additional species. The role of hormones in holocephalan reproduction is also largely unknown and deserves attention; however, information on this group will certainly be advanced in the near future because of the recent survey-sequencing of the holocephalan elephant shark (C. milii) genome. This sequencing project has already provided important

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information on evolutionary relationships and tissue distributions of several reproductive-related hormone families (Gwee et al., 2009; Larsson et al., 2009). In addition, there is a need for more experimental and functional studies to define the role of certain hormones during reproductive events, but this first requires fundamental information on the biochemical and physiological mechanisms employed by different species. It also will be important to further our understanding of the distribution of hormone receptors in both male and female chondrichthyans in order to fully appreciate the functions of reproductive hormones in these fishes and the evolution of these systems in all vertebrates.

ACKNOWLEDGEMENTS We thank members of the Tricas Lab and Gelsleichter Lab, Charles A. Manire, and staff of Mote Marine Laboratory for years of stimulating discussions on elasmobranch reproductive endocrinology and behavior. James Gelsleichter acknowledges the University of North Florida for providing the time and resources needed to prepare this chapter.

ABBREVIATIONS 11-KT 17b-HSD 3b-HSD 5-HT ABP AR AVT Cd CORT CT CYP19 DHT E2 ER FSH GnRH GSI GTH HPA HPG ir LH OC P4 P450aro POA PR T T3 T4 TN Vtg

11-ketotestosterone 17b-hydroxysteroid dehydrogenase 3b-hydroxysteroid dehydrogenase Serotonin Androgen-binding protein Androgen receptor Arginine vasotocin Cadmium Corticosterone Calcitonin See P450aro 5a-dihydrotestosterone 17b-estradiol Estrogen receptor Follicle-stimulating hormone Gonadotropin-releasing hormone Gonadosomatic index Gonadotropin Hypothalamicepituitaryeadrenal Hypothalamicepituitaryegonadal Immunoreactive Luteinizing hormone Organochlorine pesticide Progesterone Aromatase enzyme Preoptic area Progesterone receptor Testosterone Triiodothyronine Thyroxine Terminal nerve Vitellogenin

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Chapter 12

Hormones and Reproduction of Sarcopterygian Fishes Jean M.P. Joss Macquarie University, Sydney, NSW, Australia

SUMMARY There is very little reported data concerning reproduction and hormones of reproduction for the two living groups of sarcopterygian fishes, the coelacanths and the lungfishes. In general terms, coelacanths are viviparous and lungfishes are oviparous. Both are large fishes and very long-lived. They inhabit the warmer latitudes of the southern hemisphere, the coelacanths being confined to the more even conditions of the deep oceans and the lungfishes to the very variable conditions of subtropical freshwater streams and rivers. Their differing modes of reproduction are described in this chapter and some comparisons with other vertebrates are made.

1. INTRODUCTION Which are the sarcopterygian fishes? The bony fishes, class Osteichthyes, are divided into the actinopterygian or rayfinned fishes and the sarcopterygian or lobe-finned fishes (Figure 12.1). There are very few of this latter group alive today and yet they were a conspicuous part of the fish fauna during the Devonian, the ‘great age of fishes.’ The tetrapodomorph sarcopterygians (osteolepiforms, rhizodonts, and elpistostegalians) were the direct relatives of the ancestral tetrapods (e.g., Acanthostega) and they were also the most abundant sarcopterygian fishes during the Devonian, closely followed by the dipnoans (lungfishes). It is this last group along with the coelacanths that are the only sarcopterygian fishes alive today. Controversy has surrounded these two groups ever since a living coelacanth was first discovered in 1938dwhich of these two living sarcopterygian fishes is most closely related to the tetrapods? At first the coelacanth was hailed as a ‘living fossil’da link between fishes and tetrapodsdbut, as information about living coelacanths gradually accumulated, it was realized that morphologically the coelacanth does not appear to be the missing link after all (Forey, 1988). In 1981, Rosen, Forey, Gardiner, and Patterson reinstated lungfishes as the


actual ancestors of the tetrapods to the exclusion of all other fossil groups. This work initiated a very vigorous debate at a time when molecular phylogenetics was beginning to be used to unravel phylogenetic relationships among many living groups of organisms, including those at the fishetetrapod transition. At first, there appeared to be support for both lungfishes and coelacanths (Joss, 1993). Gradually, this changed to equal support for a lungfish/ tetrapod clade and a lungfish/coelacanth clade (Zardoya et al., 1998; Takezaki, Figueroa, Zaleska-Rutezynska, Takahata, & Klein, 2004) until most recently support turned toward the lungfishes, excluding coelacanths, as the closest living group to the ancestral tetrapods (Brinkman, Venkatesh, Brenner, & Meyer, 2004; Hallstrom & Janke, 2008). This has increased interest in lungfishes as the animals of choice for studying the fishetetrapod transition. Unfortunately, reproduction and the hormones of reproduction have not been the subject of much of this interest to date, primarily because of their relative inaccessibility to science. Neoceratodus is the most accessible but it is large in size (> 5 kg) and age (~20 years) at maturity and has so far proved intractable to usual aquaculture techniques. All of the living sarcopterygian fishes are found in the warmer latitudes of the southern hemisphere. The two known living species of coelacanths inhabit the deep oceans (90e200 m deep) off the east coast of Africa and the islands of Indonesia. As a result of the difficulty of obtaining these fish, they only recently have been discovered by scientists (the West Indian Ocean coelacanth (Latimeria chalumnae) in 1938 off the east coast of Africa and the Indonesian coelacanth (Latimeria menadoensis) in 1997 off the islands of Sulawesi in the archipelago of Indonesia (Erdmann, Caldwell, & Moosa, 1998). Living lungfishes are all found in relatively shallow freshwater streams, lakes, and rivers. Lepidosiren paradoxa of South America was the first lungfish to be discovered, in 1831 (Natterer, 1837). The South American lungfish together 239

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RAY-FINNED FISHES

LUNGFISHES COELACANTHS

TETRAPODS (AMPHIBIANS REPTILES, BIRDS MAMMALS)

FIGURE 12.1 Phylogentic relationships between sarcopterygians, actinopterygians, other fishes, and tetrapods.

LOBE-FINNED FISHES

CARTILAGINOUS FISHES

BONY FISHES

JAWED FISHES

with the four African species of Protopterus (West African lungfish (Protopterus annectens), marbled lungfish (Protopterus aethiopicus), East African lungfish (Protopterus amphibious), and the slender or spotted lungfish (Protopterus dolloi)) belong to the family Lepidosirenidae, which is of more recent origin. The more ancient family of Ceratodontidae is represented today by the single species, Neoceratodus forsteri, found in only a few small coastal rivers in southeast Queensland, Australia. Although the lungfishes are more accessible for research than the coelacanths, little more is known about their reproduction and related hormones than is known for the coelacanths. This chapter will review what is known about the reproduction and hormones of both groups and will consider what we may learn from sarcopterygian fishes about their group’s role as an ancestor to amphibians and amniote vertebrates.

2. COELACANTHS There has been almost nothing published on the reproduction of coelacanths since 1991, and very little on their hormones. The two review chapters written by Wourms, Atz, and Stribling, and Balon in the Biology of Latimeria chalumnae and Evolution of Coelacanths in 1991 provide coverage of what is known about coelacanth reproduction. Coelacanths are viviparous, employing yolk sac placentae for nourishment, excretion and/or gaseous exchange etc. of the developing young, not dissimilarly to most modern elasmobranchs. It was not until 1975 that Latimeria was found to be viviparous, recorded as ovoviviparous (Smith, Rand, Schaeffer, & Atz, 1975). A formalin-fixed specimen held by the American Museum of Natural History was found to contain five pups in her right oviduct/uterus (Atz, 1976). Each embryo was contained within its own ‘compartment’ with its yolk sac closely adhering to a very vascularized

region of the compartmental wall of the maternal uterus. Wourms et al. (1991) concluded from his study that nourishment of coelacanths developing within the maternal reproductive tract would have to include more than the yolk available to it from its yolk sac. Indeed, the yolk sacs of latestage pups were found to be flaccid, containing little yolk. This is indicative of some form of matrotrophy (maternal provision of nutrition for the developing pup beyond that of the yolk). Some lecithotrophy (absorption of ovulated eggs) is also suspected during earlier stages of development because of the much larger numbers of eggs being ovulated than could possibly develop within an oviduct. Gestation in Latimeria has been estimated to last 13e15 months, the pups being fully developed and able to fend for themselves at birth. Details of reproductive behavior and internal fertilization have not been recorded. None of the hormones of reproduction have been reported for coelacanths. The only anterior pituitary peptides to have been studied are those of the pro-opiomelanocortin group (Takahashi, Yasuda, Sullivan, & Kawauchi, 2003). The gonadotropins (GTHs) have not been reported and there are no descriptions of gonads and gonadal hormones to the knowledge of this writer.

3. LUNGFISHES Our knowledge about lungfish reproduction is rather greater than that for coelacanths, but is still scant in comparison with what is known about teleosts or even chondrichthyan fishes. Much remains to be learned. While not as large as coelacanths, lungfishes are still quite big fishes (adult Neoceratodus in excess of 20 kg are not rare in their native habitat). They therefore present difficulties for aquaculture techniques, in contrast to most freshwater teleosts. Nevertheless, there is at least one captive breeding program associated with a research institution in Australia. This

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Hormones and Reproduction of Sarcopterygian Fishes

facility, at Macquarie University in Sydney, coupled with careful observations of lungfishes in the wild, has provided the bulk of the information presented in this chapter. Lungfishes, unlike coelacanths, are oviparous. They produce large gametes, eggs, and sperm, housing very large nuclei apparently made necessary by their very large genomes (Rock, Eldridge, Champion, Johnston, & Joss, 1996; Gregory, 2002), and, in the case of the eggs, yolk equivalent to that produced in amphibian or primitive actinopterygian eggs. Neoceratodus may spawn several times during the late winterespring months, depending on the condition of the fish and the prevailing weather conditions over the spawning season. This and other aspects of reproduction will be discussed in the following sections. For more detailed information on the development of lungfishes, see Joss (2009).

3.1. Spawning Neoceratodus grow slowly, reaching maturity at approximately 20 years of age. Some males mature at a smaller size than females, but all fish over 6 kg in weight tend to be mature. Their gonads commence recrudescence in midautumn (fall), gradually reaching full maturity by the end of winter, when they represent ~10% of the weight of each fish. However, the gametes are large, as mentioned above, and are only released under appropriate conditions. At the end of the late winter/spring spawning season, the gonads regress. During quiescence, the testes contain no mature sperm and the ovaries present atretic oocytes, in which the yolk is being ingested by macrophages (Figure 12.2).

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The small coastal river systems that Neoceratodus naturally inhabits are shallow and slow-moving most of the time but are subject to flooding every eight to nine years, generally during the autumn. Spawning is an annual event provided that there has been adequate nourishment available over the previous year and that the shallow riffles (Figure 12.3) favored for spawning are available and wellvegetated. Adult fish tend to gather in close proximity to one of these riffles, spending the daylight hours in the main river flow area. Approaching dusk, they move into the shallow riffles and can be seen very clearly as they pair off and commence their rather casual and yet elaborate courtship rituals (Grigg, 1965). The female deposits one, or perhaps two, eggs at a time and these are fertilized by the male before the pair move off together to another place within the large riffle and repeat the process. There are no exact data but it is estimated that on a good night a pair may be responsible for perhaps as many as 200 fertilized eggs (not a large number in comparison to many teleost fishes). Following fertilization, the outer membrane of the egg becomes ‘sticky’ and adheres to the vegetation, quickly attracting particulate matter in the water that renders it well-camouflaged until the little lungfish hatches some three to five weeks later, depending on temperature. The hatchlings may remain in the same area for as long as six months to two years or more. Being very cryptic, they were at first thought to have died, which prompted a period of relocations of adult fish into several other lakes and river systems in southeast Queensland in the earlier part of the 20th century (Arthington, 2008). Some of these populations may have survived but proper studies of lungfish populations in southeast Queensland are in their infancy. Details of protopterid spawning behavior are poorly known as it takes place within ‘nests’ constructed in slowmoving, shallow sections of rivers in central Africa (Protopterus spp.) or South America (L. paradoxa).

3.2. Parental Care of Young

FIGURE 12.2 Section of a mature ovary of the Australian lungfish (Neoceratodus forsteri) in the summer, undergoing regression. Portions of several large yolky oocytes are shown along with much smaller, unyolked, immature oocytes. The yolk is no longer evenly distributed throughout the regressing (atretic) oocytes. Many large macrophages are clustered alongside the oocytes, darkly pigmented following ingestion of yolk from unovulated follicles stained with Haematoxylin and eosin. Magnification 800. See color plate section.

Neoceratodus provides no further care for the eggs/hatchlings. The lepidosirenid lungfishes, however, construct nests and the male parent may guard and aerate the nest until the hatchlings have developed their lungs, shed their external gills, and left the nest (Johnels & Svensson, 1954; Bouillon, 1961; Greenwood, 1986). In order to enrich the oxygen in the nest, male Lepidosiren develop structures on the pelvic fins that may release additional oxygen into the water in the nest (Bruton, 1998).

3.3. Hormones Involved in Reproduction There is very little published data on the hormones of reproduction in lungfishes. There are, however, several older studies on the cell types in the pituitaries of lungfishes

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FIGURE 12.3 Shallow riffle on the Burnett River, southeast Queensland, Australia, used for spawning by Neoceratodus. To the far right is the main river to which the lungfish return after each spawning bout. The figure in the centre is Greg Joss trawling for lungfish eggs, following a spawning bout the previous evening.

and the coelacanths, using heterologous antibodies for gonadotrope identification. The anterior pituitary of the living coelacanths is elongated rostrally into a separate lobe housing the gonadotropes (Lagios, 1975), as also occurs in elasmobranchs, whereas that of lungfishes is morphologically arranged like the pituitaries of amphibians, the lepidosirenid pituitaries more so than that of Neoceratodus (Joss, Beshaw, Williamson, Trimble, & Dores, 1990). What has been published about the hormones themselves tends to focus on sequencing of GTHs rather than on the physiological roles of pituitary and gonadal hormones. There is one paper by Joss, Edwards, and Kime (1996) that identifies the dominant androgen of male Neoceratodus as testosterone (T), which when seasonally measured in the circulation is highest during the spawning season (Figure 12.4) and lowest in summer. This paper also describes the production of soluble conjugated androgens by males during the spawning season. Pheromones of lungfishes is a topic about which very little has been published. However, unpublished observations of increased ventilatory frequency in response to androstenedione (AND) and T glucuronides introduced into the water, coupled with the above published demonstration of conjugated androgens (AND and T glucuronides) being produced only during the spawning season, provide circumstantial evidence that spawning events in wild populations may be synchronized by pheromones released from an alpha male in each subpopulation that returns to and uses a particular spawning site along the river each year. Comparable data on steroid hormones of reproduction in females have not been published. However, the steroidal response to the hypothalamic gonadotropin-releasing hormone (GnRH) has been published (Joss, King, & Millar, 1994), showing that, in females, 17b-estradiol (E2) can be measured in the blood two minutes after a cardiac injection

of 1.5 mg of mammalian GnRH (mGnRH) in early or late spring but not in early autumn (Figure 12.5). These data obtained for both males and females suggest that either the gonads or the pituitary gonadotropes or both are refractory for a period following the spawning season. Mammalian GnRH and chicken GnRH-II are present in lungfishes (Neoceratodus (Joss et al., 1994); Protopterus (Vallerino, Trabucchi, & Vaudry, 1998)).

FIGURE 12.4 Plasma testosterone (ng/ml) in mature male lungfish (Neoceratodus forsteri) from the Mary River (Queensland, Australia) over the years 1985e1990. The vertical lines through the means extend one standard error of mean (SEM) in each direction. The numbers in brackets refer to the sample size for each point. Reprinted from Joss, Edwards, and Kime (1996), with permission.

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4. CONCLUDING REMARKS

FIGURE 12.5 (a) Changes in serum 17b-estradiol (E2) in female lungfish (Neoceratodus forsteri) in response to a single cardiac injection of 1.5 mg/kg mammalian GnRH (mGnRH ¼ GnRH-I) administered in early spring (-), late spring (,), and early autumn (A). Values represent means from two lungfish from each of the sampling times. (b) Changes in serum testosterone (T) in male lungfish in response to a single cardiac injection of 1.5 mg/kg mGnRH administered in early spring (three fish) (-), late spring (four fish) (,), and early autumn (three fish) (A). Adapted from Joss, King, and Millar (1994).

Pituitary GTHs have been sequenced by cDNA techniques for Neoceratodus (Arai, Kubokawa, Ishii, & Joss, 1998; Querat et al., 2004). The physiology of these hormones has not been investigated for lungfish but the sequence similarity is sufficiently like that of amphibians and mammals to suggest that the hypothalamicepituitaryegonadal axis should operate in the manner described for these tetrapods. Namely, GnRH from the hypothalamus, under feedback control from gonadal steroid hormones and the central nervous system (CNS) responses to diurnal and seasonal cues, stimulates the release of GTHs from the pituitary. The GTHs act on the gonads to release androgens synthesized by the testes and estrogens by the ovaries, which in turn control the maturation of gametes, including the sequestering of vitellogenin, produced in the liver, by growing oocytes in the ovary. Gonadotropins may control spermiation and ovulation, while steroid hormones may act on the brain to stimulate spawning behaviors. Currently, there are few data to confirm or deny whether hormones behave generally in lungfishes to control maturation and release of gametes or spawning behavior in this way.

The two living groups of sarcopterygian fishes could not be more different in their modes of reproduction. Coelacanth reproduction is more like that of advanced elasmobranches (see Chapter 11, this volume) and lungfish reproduction like that of living amphibians (see Volume 2). These affinities are also reflected in the morphology of the anterior pituitary of the two groups, as mentioned above. These disparate modes of reproduction can tell us very little, and nothing of certainty at this stage of their study, about their evolutionary relationship with each other or that of either of them with tetrapods or other fish groups. Probably the most that can be said is that both groupsdlungfishes and coelacanthsdare very ancient fishes, lungfishes having closer affinities with later sarcopterygians and coelacanths possibly with cartilaginous fishes, all of which were evolving at the Silurian/Devonian border. The very recent discovery (Long, Trinajstic, & Johanson, 2009) that some Devonian placoderm fishes were viviparous, like the coelacanths and elasmobranches today, offers some intriguing alternatives to the current thought that coelacanths are the closest relatives of lungfishes. We cannot leave the discussion of sarcopterygian reproduction without considering field studies urgently needed on Neoceratodus to inform plans for their survival in the face of Australia’s increasing water problem. The few little coastal rivers close to Brisbane that the lungfish inhabit are under increasing threat of damming to supply much-needed water to Brisbane’s growing population and for irrigation for crops such as sugar cane. Both behavioral and pheromonal studies are needed along with the population studies that will inform us unequivocally of the effects of dams on lungfish spawning and recruitment. This particular species of lungfish has survived unchanged for at least 100 million years. Can it also survive this most recent threat?

ABBREVIATIONS AND CNS E2 GnRH GTH T

Androstenedione Central nervous system 17b-estradiol Gonadotropin-releasing hormone Gonadotropin Testosterone

REFERENCES Arai, Y., Kubokawa, K., Ishii, S., & Joss, J. M. P. (1998). Cloning cDNA encoding the common alpha subunit precursor molecule of pituitary glycoprotein hormones in the Australian lungfish, Neoceratodus forsteri. Gen. Comp. Endocrinol., 110, 109e117.

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Arthington, A. H. (2008). Australian lungfish, Neoceratodus forsteri, threatened by a new dam. Environ. Biol. Fish., 10, 1007/s10641-0089414-y (published early on line September 2008). Atz, J. W. (1976). Latimeria babies are born, not hatched. Underwater Naturalist, 9, 4e7. Balon, E. K. (1991). Probable evolution of the coelacanth’s reproductive style: lecithotrophy and orally feeding embryos in cichlid fishes and in Latimeria chalumnae. Environ. Biol. Fish., 32, 249e265. Brinkmann, H., Venkatesh, B., Brenner, S., & Meyer, A. (2004). Nuclear protein-coding genes support lungfish and not the coelacanth as the closest living relatives of land vertebrates. Proc. Nat. Acad. Sci., 101, 4900e4905. Bruton, M. N. (1998). Lungfishes and coelacanth. In J. R. Paxton, & W. N. Eschmeyer (Eds.), Encyclopedia of Fishes (pp. 70e74). San Diego, CA: Academic Press. Erdmann, M. V., Caldwell, R. L., & Moosa, M. K. (1998). Indonesian ‘king of the sea’ discovered. Nature, 395, 335. Forey, P. L. (1988). Golden jubilee for the coelacanth, Latimeria chalumnae. Nature, 336, 727e732. Gregory, T. R. (2002). Genome size and developmental complexity. Genetica, 115, 131e146. Grigg, G. C. (1965). Spawning behaviour in the Queensland lungfish, Neoceratodus forsteri. Australian Natural History, 15, 75. Joss, J. M. P. (2009). Sarcopteygian fish. In Y. Kunz-Ramsay (Ed.), Development of Non-Teleostean Fishes. New Hampshire: Science Publishers. Joss, J. M. P. (1993). Dipnoan evolution: molecular studies. Proceedings of the Zoological Society of Calcutta, Haldane Commemorative Volume, 93e98. Joss, J. M. P., Beshaw, M., Williamson, S., Trimble, J., & Dores, R. M. (1990). The adenohypophysis of the Australian lungfish, Neoceratodus forsteridan immunocytological study. Gen. Comp. Endocrinol., 80, 274e287. Joss, J. M. P., Edwards, A., & Kime, D. E. (1996). In-vitro biosynthesis of androgens in the Australian lungfish, Neoceratodus forsteri. Gen. Comp. Endocrinol., 101, 256e263. Joss, J. M. P., King, J. A., & Millar, R. P. (1994). Identification of the molecular forms of and steroid hormone response to gonadotropin-releasing


hormone in the Australian lungfish, Neoceratodus forsteri. Gen. Comp. Endocrinol., 96, 392e400. Long, J. A., Trinajstic, K., & Johanson, Z. (2009). Devonian arthrodire embryos and the origin of internal fertilisation in vertebrates. Nature, 457, 1124e1127. Natterer, J. (1837). Lepidosiren paradoxa, eine neue Gattung aus der Familie der fischaenlichen Reptilien. Annal. Naturhistor. Mus., 2, 165e170, [reference obtained fromVanzolini, 1996]. Querat, B., Arai, Y., Henry, A., Akama, Y., Longhurst, T., & Joss, J. M. P. (2004). Pituitary glycoprotein hormone b subunits in the Australian lungfish and estimation of the relative evolution rate of these subunits within vertebrates. Biol. Reprod., 70, 353e363. Rock, J., Eldridge, M., Champion, A., Johnston, P., & Joss, J. M. P. (1996). Karyotype and nuclear DNA content of the Australian lungfish, Neoceratodus forsteri (Ceratodidae: Dipnoi). Cytogenet. Cell. Genet., 73, 187e189. Rosen, D. E., Forey, P. J., Gardiner, B. G., & Patterson, C. (1981). Lungfishes, tetrapods, paleontology and pleisiomorphy. Bull. Amer. Mus. Nat. Hist., 167, 159e276. Smith, C. L., Rand, C. S., Schaeffer, B., & Atz, J. W. (1975). Latimeria, the living coelacanth is ovoviviparous. Science, 190, 1105e1106. Takahashi, A., Yasuda, A., Sullivan, C., & Kawauchi, H. (2003). Identification of proopiomelanocortin-related peptides in the rostral pars distalis of the pituitary in coelacanth: evolutional implications. Gen. Comp. Endocrinol., 130, 340e349. Takezaki, N., Figueroa, F., Zaleska-Rutezynska, Z., Takahata, N., & Klein, J. (2004). The phylogenetic relationship of tetrapod, coelacanth, and lungfish revealed by the sequences of forty-four nuclear genes. Mol. Biol. Evol., 21, 1512e1524. Vallarino, M., Trabucchi, M., & Vaudry, H. (1998). Neuropeptides in the lungfish brain: phylogenetic implication. Ann. NY Acad. Sci., 839, 53e59. Vanzolini, P. E. (1996). A contribuiçaõ zoolo´gica dos primeiros naturalistas viajantes no Brasil. Rev. USP, 30, 190e238. Wourms, J. P., Atz, J. W., & Stribling, D. (1991). Viviparity and the maternaleembryonic relationship in the coelacanth Latimeria chalumnae. Environ. Biol. Fish., 32, 225e248.

Chapter 13

Endocrine-active Chemicals (EACs) in Fishes Alan Milan Vajda* and David O. Norrisy *

University of Colorado at Denver, Denver, CO, USA, y University of Colorado at Boulder, Boulder, CO, USA

SUMMARY Vertebrate endocrine systems coordinate complex developmental, physiological, and behavioral events to mediate lifehistory tradeoffs and to optimize fitness. Information from diverse biotic and abiotic factors including temperature, photoperiod, chemicals, and the social environment are integrated and transduced into developmental and physiological responses via neuroendocrine regulatory networks. Exogenous endocrineactive chemicals (EACs) from diverse structural classes, of natural or anthropogenic origin, can interact with these regulatory networks and disrupt, mimic, or otherwise interfere with endocrine-mediated regulatory processes. Effects of such perturbations range from transient, reversible deviations from homeostasis to the irreversible disruption of reproductive differentiation, heritable epigenetic impacts, or local population extinctions. Fishes and other aquatic vertebrates appear to be especially susceptible to the adverse effects of EAC exposure as the aquatic environment appears to be a sink for natural and anthropogenic EACs.

1. INTRODUCTION The fishes, and especially the teleost fishes, are the most abundant vertebrate group on Earth in terms of both number of species and total biomass. Further, as the oldest group of vertebrates, they exhibit the greatest diversity in reproductive modes, making them obvious targets for scientific studies of ecology, physiology, and evolution. This volume has documented much of what we know about fish reproduction and its regulation by hormones produced by the endocrine system and how secretion of these hormones is influenced by environmental factors operating through neuroendocrine mechanisms of the hypothalamicepituitaryegonadal (HPG), hypothalamicepituitaryeadrenal/interrenal (HPA/HPI), and hypothalamusepituitaryethyroid (HPT) axes. All vertebrate neuroendocrine systems coordinate complex developmental, physiological, and behavioral events to optimize fitness. Information received from diverse biotic and abiotic factors including temperature, photoperiod, natural chemicals, and


the social environment are integrated and transduced into developmental and physiological responses via these neuroendocrine mechanisms. The HPG axis is the primary axis regulating reproduction, involving the production in the hypothalamus of gonadotropin-releasing hormone (GnRH), which stimulates secretion of the gonadotropins (GTHs) from the pituitary: follicle-stimulating hormone (FSH) and luteinizing hormone (LH) (for details see Chapter 2, this volume). Cooperatively, these GTHs regulate production of gametes and synthesis of gonadal hormones including androgens and estrogens. The major androgens produced in fishes are testosterone (T) and 11-ketotestosterone (11-KT), and the major estrogen is 17b-estradiol (E2). Androgens and estrogens produce their effects in a variety of target tissues by binding to androgen receptors (ARs) and estrogen receptors (ERs), respectively. Despite the diversity of fishes and their reproductive modes, the hormones involved and the cellular and genetic mechanisms through which they operate are fundamentally the same as those described in the other volumes of this series that focus on hormones and reproduction in amphibians, reptiles, birds, and mammals. Modern fishes are important for a variety of commercial purposes including human food, and globally we consume billions of tons of fishes annually. Consequently, the health and reproductive status of fishes should be of primary concern for practical reasons as well as for their scientific value. In addition to being a major consumer of fishes, the increasing global human population also is the major source of chemical wastes that find their way into freshwater ecosystems, through which they are transported to estuaries and coastal regions that represent the most critical biological components of marine ecosystems. Many of these chemicals have been shown to disrupt endocrine systems by either mimicking natural hormones, blocking hormone actions, or altering secretion, mechanisms of action, or rates of metabolism of endogenous hormones. 245

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These exogenous chemicals, whether they may be of natural or anthropogenic origin, may be termed endocrineactive chemicals (EACs), and they can interact with regulatory networks, and disrupt, mimic, or otherwise interfere with endocrine-mediated regulatory processes. Effects of such perturbations range from transient, reversible deviations from homeostasis to irreversible disruption of reproductive differentiation, heritable epigenetic impacts, or local population extinctions. As the aquatic environment acts as a sink for natural and anthropogenic EACs, fishes and other aquatic vertebrates appear to be especially susceptible to the adverse effects of EAC exposure. Consequently, this chapter will focus on water as the vector for bringing anthropogenic EACs to wild populations of fishes.

1.1. Endocrine Disruption in Fishes Although vertebrates have been exposed to EACs and have adapted to their presence over millions of years, the acute discharge of thousands of diverse and persistent EACs from the activities of industrialized humans is unparalleled in their evolutionary history (reviewed by Vajda & Norris, 2006). Industrialized societies have synthesized and released more than 80 000 chemicals into the environment in the last 50e70 years (Curtis & Skaar, 2002) and have dramatically altered the distribution and aquatic occurrence of endocrine-active metals, dioxins, phytochemicals, and biogenic steroids. This represents a dramatic radiation of potentially disruptive signal molecules that rivals major paleoecotoxicological events such as the redistribution of crustal heavy metals following extraterrestrial collisions (Herkovits, 2001) or the diversification of bioactive endocrine-active phytochemicals that accompanied the angiosperm radiation of 200 million years ago (reviewed in Stafford, 2000). One of the most complex and heterogeneous functional classes of these synthetic compounds includes those that act upon the neural and endocrine regulation of reproduction in vertebrates, altering the expression of life-history phenotypes. In anthropogenic settings, even evolutionarily ancient biogenic signaling molecules can destabilize signaling networks through chronic out-of-context signaling (reviewed in McLachlan, 2001). Human settlements and industrial livestock operations are responsible for the pointsource discharge of biologically relevant concentrations of biogenic steroids into the aquatic environment. These molecules are structurally and functionally similar to those endogenous steroid regulators of signaling pathways central to vertebrate fitness. Exogenously derived biogenic estrogens can interact directly with estrogenic regulatory systems in vertebrates and are interpreted as molecules with high information content. The high concentrations of EACs associated with anthropogenic municipal, agricultural, or


industrial effluents often yield adverse effects in exposed organisms, including plants (Fox, Starcevic, Kow, Burrow, & McLachlan, 2001), invertebrates (Oberdorstor et al., 2001), and vertebrates (reviewed in McLachlan, 2001). Endocrine-active chemicals from diverse sources have been identified from the surface waters of North America (Kolpin et al., 2002), Australia (Williams et al., 2007), Asia (Lei, Huang, Zhou, Wang, & Want, 2009), Africa (Barnhoorn, Bornman, Pieterse, & Van Vuren, 2004), and Europe (Allen et al., 1999; Adler, Steger-Hartmann, & Kalbfus, 2001). Evidence of reproductive disruption has been observed in freshwater, saltwater, and anadromous fishes (Segner, 2005) inhabiting watersheds impacted by urban (Jobling, Nolan, Tyler, Brighty, & Sumpter, 1998; Woodling, Lopez, Maldonado, Norris, & Vajda, 2006; Vajda et al., 2008), agricultural (Orlando et al., 2004), and industrial activities (Munkittrick, McMaster, McCarthy, Servos, & Van der Kraak, 1998) as well as from pristine habitats with relatively little direct human impact (Schwindt et al., 2009). Some EACs are natural components of aquatic ecosystems, and some naturally occurring EACs, such as some endocrine-active phytochemicals (EAPs), may play a signaling role in normal vertebrate physiology (reviewed in Vajda & Norris, 2006). However, rapidly expanding human populations and activities have profoundly altered the occurrence and concentration of EACs in aquatic ecosystems. The signaling noise caused by the introduction of anthropogenic EACs into aquatic environments has become an intrinsic signaling component of many aquatic environments. A corruption of environmental signal integrity by noise from anthropogenic signaling may be another hallmark of the Anthropocene Age (Meybeck, 2003; 2004).

2. MECHANISMS OF ENDOCRINE-ACTIVE CHEMICAL (EAC) SIGNALING Endocrine-active chemicals from diverse structural classes including steroids, organochlorines, dioxins, polychlorinated biphenyls (PCBs), alkylphenolic surfactants, flavonoids, and other phytochemicals, inorganic anions, and metals can signal via various central and peripheral components of the fish reproductive axis (Pait & Nelson, 2003; Kiparissis et al., 2003; Guillete & Edwards, 2005; Vetillard & Bailhache, 2005; Bistodeau et al., 2006). Exposure of fishes to EACs can potentially disrupt any and every protein-mediated interaction underlying the endocrine- and neuroendocrine-mediated regulation of reproductive differentiation, development, behavior, and function. Endocrine-active chemicals modulate fish reproduction by interfering with the production (Ankley et al., 2002), release, transport (Morgado, Hamers, Van der Ven, & Power, 2007), metabolism, binding, action, or

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elimination of endogenous hormones (Kavlock et al., 1996), with life-stage-specific consequences. Many EACs signal through multiple life stages via tissue-specific pathways resulting in complex outcomes, especially from exposure to complex environmental mixtures. Specificity in ligand recognition and capacity to discriminate signal from noise are key components of the integrity of signaling networks. Vertebrate endocrine signaling networks have been challenged, and perhaps shaped, by exogenously derived EACs from dietary and environmental sources throughout their evolution. In modern times, however, vertebrate endocrine systems are encountering novel synthetic chemicals as well as familiar biogenic EACs in novel signaling contexts.

2.1. Estrogenic Signaling Pathways Nuclear receptors are an evolutionarily ancient superfamily of ligand-activated transcription factors that bind diverse dietary, environmental, and endogenous ligands. Nuclear receptor-mediated signaling regulates life-history traits through actions on diverse gene networks. The best-characterized actions of EACs on fish reproduction are mediated via members of the steroid receptor subfamily of nuclear receptors. The ER is the most ancient member of the steroid receptor subfamily (Thornton, 2001); receptors for the other steroids are hypothesized to have arisen following duplication and diversification of the ancestral ER. Although its native endogenous ligand, E2, is the ultimate product of steroid biosynthesis in vertebrates, it appears to have been the first steroid to signal via nuclear receptor-mediated pathways (Thornton, 2001). Signaling through steroid-signaling pathways is an evolutionarily ancient means of communication within and between organisms; these pathways regulate reproduction in some plants (Geuns, 1978; Grunwald, 1980), many invertebrates (S. Atkinson & M. Atkinson, 1992; Baldwin & LeBlanc, 1994; Tarrant, S. Atkinson, & M. Atkinson, 1999; LeBlanc & McLachlan, 2000), and all vertebrates. Despite differences in gonadal plasticity and the relative contributions of genotype and environment to gonadal sex specification, steroid hormones are central to developmental organization and activational regulation of reproduction in all vertebrates (Guillette, Cram, Rooney, & Pickford, 1995).

2.1.1. Estrogen receptors (ERs) The ER in teleost fishes exists as multiple subtypes (Hawkins et al., 2001). Each ER subtype is encoded by a different gene and differs in its tissue distribution and ligand preference (Mosselman, Polman, & Dijkema, 1996; Kuiper et al., 1997). In addition to reproductive tissues, ERs are found in the liver, gut, central and peripheral nervous

systems, blood vessels, skin, and other tissues (Kuiper et al., 1997). Through interactions with ERs within central and peripheral target tissues, estrogens regulate growth, differentiation, maturation, and function of reproductive and nonreproductive tissues. Modulation of estrogendependent genes in target tissues occurs within narrow E2 concentrations in the pico- to femto-molar range (Welshons et al., 2003). Small fluctuations in local concentrations of E2 during critical developmental stages can induce dramatic changes in ER-modulated gene networks, yielding outcomes with adverse fitness consequences including inappropriate sexual differentiation of the gonad or brain (Welshons et al., 2003; see also Chapter 1, this volume). The required precision of dosage, timing, and coordination of gene expression add to the susceptibility of various stages of sex differentiation to disruption by external influences. Such disruption of the hormonal milieu can result from numerous stressors or environmental agents. Among the most insidious and ancient challenges to the integrity of endocrine signaling networks is the inappropriate interaction with exogenously derived EACs.

2.1.2. Estrogen receptor (ER) ligand promiscuity The ER binds several structurally diverse classes of chemicals including steroids, metals, flavonoids, fatty acids, and peptides (Veeneman, 2005). This promiscuity appears to be unique among nuclear receptors and has been partially attributed to the size of the ER ligand-binding pocket, which is almost twice the volume of E2 (Brzozowsi et al., 1997; Elsby et al., 2000). This apparent ligand promiscuity may reflect an adaptation to permit the modulation of reproduction and other life-history traits by diverse dietary and environmental ligands. Alternatively, promiscuity in ligand recognition may reflect an evolutionary constraint imposed by the contribution of the ER to fitness and high connectivity to other signaling systems (Fraser, Hirsh, Steinmetz, Scharfe, & Feldman, 2002). Promiscuity in ligand recognition may decrease the capacity of estrogenic signaling networks to discriminate signal from noise in complex information environments, destabilizing signal network integrity.

2.1.3. Estrogen additivity Mixtures of estrogenic chemicals act additively, according to principles of concentration addition, for some ERdependent endpoints both in vitro (Silva, Rajapakse, & Kortenkamp, 2002) and in vivo (Thorpe et al., 2006; Brian et al., 2007). Estimates of total estrogenicity may be useful in evaluating impacts upon vitellogenesis and other phenomena primarily regulated by a single ER subtype (Thorpe et al., 2003; Brian et al., 2005; 2007). However, simple additivity-based estrogenicity estimates may be

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limiting when examining complex endpoints such as the neuroendocrine regulation of behavior or gametogenesis, where estrogens signal with compound- and pathwayspecific potency via multiple estrogenic and nonestrogenic, and genomic and nongenomic signaling pathways (Tabb & Blumberg, 2006; Thorpe et al., 2006). This critical, multiple-pathway signaling complexity is lost when waste water treatment plant (WWTP) effluent estrogen composition and concentration are reduced to the single dimension of effluent estrogenicity as determined by invitro ER-based assays (Martinovic, Denny, Schmieder, Ankley, & Sorensen, 2008b).

2.2. Androgenic Signaling Pathways The AR is a member of the same nuclear receptor family as the ER (Thornton, 2001). Although the AR is considered to have evolved more recently than the ER, the AR gene is present in all vertebrates as well as in nonvertebrate chordates (Katsu, Kubokawa, Urushitani, & Iguchi, 2010). Chonrichthyan fishes have only a single AR but two forms of ARs have been described recently in teleosts: ARa and ARb (Ogino, Katoh, Kuraku, & Tamada, 2009).

2.3. The Effect of Endocrine-active Chemicals (EACs) on Steroidogenesis and Steroid Metabolism Another pathway for EACs to alter reproduction can occur through alteration of the synthesis of endogenous androgens and estrogens. This can be accomplished through effects on the activity of key enzymes necessary for steroidogenesis in either testes (see Chapter 3, this volume) or ovaries (see Chapter 4, this volume). For example, the enzyme aromatase (P450aro) is especially of interest because it controls the conversion of T to E2 in fishes as in all vertebrates and is often examined (e.g., Fenske & Segner, 2004). Rates of metabolic inactivation of steroids through glucuronidation or other processes also can influence endogenous steroid hormone levels. Enzymes involved in these processes can also be targets for EACs and hence disrupt reproduction.

3. MULTIDIMENSIONAL MIXTURE COMPLEXITY Environmental endocrine-active effluents are complex mixtures containing multiple chemicals that may exhibit identical, overlapping, or independent modes of action (MOAs). For example, various steroidal, estrone (E1) and estriol (E3), and nonsteroidal WWTP estrogens differ widely in their relative signaling potency via genomic ER,


nongenomic ER, and non-ER pathways (Tabb & Blumberg, 2006). Notably, signaling potency by steroidal E1 and E3 and nonsteroidal alkylphenols via nongenomic ER pathways can be disproportionately high relative to their lower signaling potency via genomic ER pathways (Kochukov, Jeng, & Watson, 2008; Watson, Jeng, & Kochukov, 2008). Concentrations of estrogenic WWTP contaminants change continually and independently, and so do effluent concentrations of antiestrogens, antiandrogens/androgens, and neuroactive pharmaceuticals capable of modulating peripheral or central responses to exogenous estrogens (Martinovic et al., 2008b; Vajda et al., 2010). The composition of such mixtures is multidimensional and complex due to continuous and independent changes in the occurrence and concentration of ligands for each specific estrogenic and nonestrogenic signaling pathway.

4. CONSEQUENCES OF SPECIFIC LIFE-STAGE EXPOSURES Exposure of fishes to EACs can potentially disrupt any endocrine- or neuroendocrine-mediated interaction regulating reproductive differentiation, physiology, and behavior. The outcomes of exposure to EACs are life-stagespecific and depend upon the dosage and timing of exposure. Lower exposure dosages may require longer exposure durations to produce effects; exposure to dosages of EACs that are subthreshold for endpoints in short-term assays can yield reproductive failure and other dramatic effects if exposure durations are sufficient (Grist, Wells, Whitehouse, Brighty, & Crane, 2003; Nash et al., 2004; Parrott & Blunt, 2005; Kidd et al., 2007). For example, while short-term exposure of adult fishes to estrogenic EACs may be sufficient to induce Vtg mRNA and protein, or to disrupt reproductive behaviors (Bistodeau et al., 2007), it is unlikely to result in the induction of intersex or sexreversal. The induction of intersex, sex-reversal, or other tissue remodeling events by estrogenic EACs requires exposure to a sufficient dose during a critical window of gonadal development and is therefore most likely to result from specific early-life-stage (Van Aerle, Pounds, Hutchinson, Maddix, & Tyler, 2002), whole-lifecycle (Nash et al., 2004), or multigenerational (Kidd et al., 2007) exposures that encompass the vulnerable developmental window.

5. ORGANIZATIONAL AND ACTIVATIONAL EFFECTS OF ENDOCRINE-ACTIVE CHEMICALS (EACS) In general, exposure of organisms to exogenous EACs prior to or during tissue differentiation can result in inappropriate organizational specification of tissue fate or signaling network behavior (Beatty, 1979; Guillette et al., 1995).

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Organizational disruption of development is permanent, and in some cases heritable, and is likely mediated by epigenetic interactions (Crews & McLachlan, 2006). When exposure occurs outside of the critical window of tissue or network differentiation, endocrine-mediated processes may be perturbed activationally (Beatty, 1979; Guillette et al., 1995; Contractor, Foran, Li, & Willett, 2004). If exposed animals are in nonreproductive condition, steroid-dependent pathways may be inappropriately activated. If exposed animals are in reproductive condition, steroid-dependent reproductive endpoints may be inappropriately inhibited. Such disruption is generally reversible upon depuration (e.g., transfer of fish to clean water) (Liney, Jobling, Shears, Simpson, & Tyler, 2005). Some exposures may be restricted to a specific developmental window, and others may be continuous or intermittent throughout the life of an organism. Organizational effects typically occur during embryonic development or in larvae or neonates, whereas activational effects are typically observed in juveniles or adults. Development can be viewed as a series of critical organizational events that determine the structure and functioning of the resulting organism. These critical events generally are mediated by endogenous chemical regulators and, once complete, the results usually are permanent. Some of these critical embryonic events, including those regulated by steroid hormones, can be readily affected by EACs if they are present when a specific organizational event is in progress. Organizational events result in permanent, usually irreversible determinations. Sex determination in the hypothalamus of the mammalian brain is an example of an organized event. In contrast, activational events regulated by steroid hormones and their related EACs usually occur later in the life of an animal, e.g. changes associated with puberty or seasonal reproduction. These effects are usually reversible if the hormone or stimulatory EAC is withdrawn.

5.1. Organizational Disruption The structure and function of vertebrate reproductive physiology has been largely conserved, with broad conservation of the structure and function of the HPG axis. As in other vertebrates, steroids play central roles in both gonadogenesis and gametogenesis in fishes (see Chapters 3 and 4, this volume). A key distinction is the plasticity of gonadal development in fishes when compared with other vertebrates (Shapiro, 1992; Nakamura, Kobayashi, Chang, & Nagahama, 1998; see also Chapters 1 and 8, this volume).

5.1.1. Gonadal differentiation Mechanisms underlying sex differentiation in teleosts are diverse and labile (Francis, 1992). Genotypic sex in many

fishes can be readily influenced by environmental factors including environmental steroids (Patino, 1997; Devlin & Nagahama, 2002; Strussman & Nakamura, 2002). Partial and complete sex reversal has been induced in > 50 species of teleosts from 24 families by administration of sex steroids, their antagonists, or aromatase inhibitors (Piferrer, 2001; Devlin & Nagahama, 2002). If gonadogenesis is disrupted by sex steroid exposure of sufficient intensity or duration, complete gonadal sex reversal can be induced. Sexual differentiation in most fishes typically occurs after hatching, during early life stages. Sexual differentiation is believed to be a time when the gonad has a heightened sensitivity to disruption by EACs (Shapiro, 1992; Nakamura et al., 1998; Van Aerle et al., 2002). The differentiated gonads of some teleosts, including hermaphroditic and gonochoristic species, retain plasticity in their responsiveness to exogenous sex steroids, though higher intensity and duration of exposure may be necessary to alter gonadal sex during later life stages (Nakamura et al., 1998).

5.1.2. Gonadal intersex Gonadal intersex, or the simultaneous occurrence of ovarian and testicular tissue, is rare among adult freshwater gonochoristic teleost fishes (Strussmann & Nakamura, 2002). Although intersex occurs naturally as a transitory stage in protandric or protogynous species of fishes, in gonochoristic species intersex is not a normal part of the reproductive mode or lifecycle. In many fishes, gonadal intersex can be induced through exposure to steroidal estrogens, androgens, antiandrogens, and estrogenic alkylphenols (APs) and EAPs (see Table 13.1). In some fishes, the critical window for intersex induction is narrow and restricted to early larval stages (e.g., the fathead minnow (Pimephales promelas) (Van Aerle et al., 2002)). In other species, such as the medaka (Oryzias latipes), exposure of embryos or larvae to steroids for a sufficient duration can yield intersex (e.g., Koger, Teh, & Hinton, 2000). Gonadal intersex appears to occur in response to exposure dosages or durations of steroid exposure that are insufficient to yield complete sex reversal (see Pandian & Sheela, 1995; Strussman & Nakamura, 2002). The identification of gonadal intersex in gonochoristic teleosts is increasingly recognized as a biomarker of early life-stage (organizational) exposure to exogenous EACs. The first reports of pollution-associated gonadal intersex came from English rivers, downstream of endocrine-active WWTP effluents (Purdom et al., 1994; Harries et al., 1997; Van Aerle et al., 2001; Jobling et al., 2002a; 2002b). Intersex incidence has been correlated with WWTP effluent load and effluent estrogenicity (Jobling et al., 1998; Robinson et al., 2003). In some WWTP-effluent-contaminated rivers in the UK, intersex gonads have been found in

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TABLE 13.1 Effects of endocrine-active chemicals (EACs) on induction of gonadal intersex Species

Common name

Treatment

Reference

Cyprinus carpio

Common carp

4-tert-Pentylphenol

Gimeno et al. (1997)

Oryzias latipes

Medaka

E2E1, E2, E3, EE2EE2

Koger et al. (2000); Metcalfe et al. (2001); Seki et al. (2002)

Danio rerio

Zebrafish

EE2, 4-Nonylphenol

Hill and Janz (2003)

Oncorhynchus mykiss

Rainbow trout

E2

Krisfalusi and Nagler (2000)

Cyprinodon variegates

Sheepshead minnow

EE2

Zilloux et al. (2001)

Oryzias latipes

Medaka

TestosteroneEE2

¨ rn et al. (2006b) Koger et al. (2000); O

Sebastes schlegeli

Korean rockfish

Methyltestosterone

Lee et al. (2003)

Danio rerio

Zebrafish

Methyltestosterone

¨ rn et al. (2003) O

Oncorhynchus tshawytscha

Chinook salmon

Steroidal androgen

Piferrer et al. (1993)

Oryzias latipes

Medaka

Vinclozolin, cyproterone acetate

Kiparissis et al. (2003b)

Oryzias latipes

Medaka

4-Nonylphenol, 4-tert-octophenol

Seki et al. (2003)

Oryzias latipes

Medaka

Genistein, equol

Kiparissis et al. (2003a)

E1, estrone; E2, 17b-estradiol; E3, estriol; EE2, 17a-ethinylestradiol.

up to 100% of sampled roach (Rutilus rutilus) (Jobling et al., 1998). The identification of gonadal intersex in fishes globally (see Table 13.2) has been associated with widespread estrogenic contamination of the aquatic environment (Folmar et al., 1996; Harries et al., 1999; Folmar et al., 2001; Kolpin et al., 2002; Petrovic, Sole, Lopez de Alda, & Barcelo, 2002; Sole´, Barcelo, & Porte, 2002; Aerni et al., 2004). In most cases, the intersex condition observed in free-living fishes is histologically classified as testis-ova (Gercken & Sordyl, 2002; Vethaak et al., 2002; DeMetrio et al., 2003; Stentiford et al., 2003), where a modest number of unstimulated primary oocytes are distributed throughout an otherwise normal testis. In contrast, the ovo-testis condition is characterized as an intersex gonad in which both ovarian and testicular tissues are relatively abundant and appear to be functional. The signaling networks and molecular mechanisms underlying intersex specification, maintenance, and fate remain unknown. There is some evidence that the occurrence of gonadal intersex in free-living fishes may be a reliable indicator of potentially adverse population impacts in these fishes. Intersex fishes may exhibit reduced fertility (Jobling et al., 2002a; 2002b) or otherwise impaired reproduction (Balch, Mackenzie, & Metcalfe, 2004). Jobling et al. (2002b) found that fishes with ovo-testes are less likely to produce milt, and that the sperm produced is of lower quality, having reduced motility and reduced fertilization capacity.

Laboratory studies have demonstrated that the concentrations of E2, 17a-ethinylestradiol (EE2), and alkylphenols found in some aquatic environments that receive domestic effluents are sufficient to induce most of the feminizing effects reported in caged and wild fishes (Robinson et al., 2003; Thorpe et al., 2003). However, laboratory efforts to induce gonadal intersex in larval fish through controlled exposure to actual WWTP effluents have been largely unsuccessful. Rodgers-Gray et al. (2001) exposed larval roach (R. rutilus) to an endocrineactive effluent where a high incidence of intersex roach had been identified (Jobling et al., 1998). Exposure of early-life-stage roach to this WWTP effluent feminized reproductive ducts but did not induce gonadal intersex (Rodgers-Gray et al., 2001). Subsequent depuration of exposed fish in clean water did not reduce the incidence of feminized ducts (Rodgers-Gray et al., 2001). In ¨ rn, Svensen, Viktor, Holbech, and Norrgren contrast, O (2006) produced ovo-testes in zebrafish (Danio rerio) by exposing them to a 50% dilution of bleached kraft paper mill effluent (BKME) for 30 days. They also observed testis-ova in zebrafish similarly exposed to a 10% BKME solution. Although laboratory attempts to induce gonadal intersex through exposure of larval fish to WWTP effluents have not been successful, the presence of gonadal intersex in free-living gonochoristic teleosts is likely induced by exposures of precise timing, dosage, and duration to exogenous EACs during critical early lifestages.

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TABLE 13.2 The identification of gonadal intersex in wild fishes globally Species

Common name

Reference

Abramus abrama

Freshwater bream

Vethaak et al. (2002)

Barbus plebejus

Barbel

Vigano et al. (2001)

Catostomus commersoni

White sucker

Woodling et al. (2006); Vajda et al. (2008)

Clarias gariepinus

Sharptooth catfish

Barnhoorn et al. (2004)

Coregonus clupeaformis

Lake whitefish

Mikaelian et al. (2002)

Gasterosteus aculeatus

Three-spine stickleback

Gercken and Sordyl (2002)

Micropterus dolomieu

Small mouth bass

Hinck et al. (2009)

Micropterus salmoides

Large mouth bass

Hinck et al. (2009)

Morone americana

White perch

Kavanagh et al. (2004)

Perca fluviatilis

Perch

Gercken and Sordyl (2002)

Platichthys flesus

European flounder

Minier et al. (2000); Simpson et al. (2000); Vethaak et al. (2002); Stentiford et al. (2003); Bateman et al. (2004); Kirby et al. (2004)

Pleuronectes yokohamae

Marbled flounder

Hashimoto et al. (2000)

Rutilus rutilus

Roach

Jobling et al., 2000a; 2002b; Bjerregaard et al., 2006

Scaphirhynchus platorynchus

Shovelnose sturgeon

Harshbarger et al., 2000

Zoarces viviparus

Eelpout

Gercken and Sordyl (2002); Stentiford et al. (2003)

5.2. Activational Disruption Activational actions of steroids are well-documented among vertebrates. We discuss three examples here that are closely linked to effects of EACs seen in wild fishes.

5.2.1. Vitellogenin (Vtg) induction in male fishes Vitellogenin (Vtg) is a phospholipoglycoprotein normally secreted by the liver of female oviparous vertebrates under stimulation of estrogens (Polzonetti-Magni, Mosconi,

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Soverchia, Kikuyama, & Carnevale, 2004). This protein is incorporated into growing oocytes by the ovary under the stimulation of LH and is converted into yolk protein. Sexually immature female and male fishes do not normally produce substantial quantities of Vtg. However, the hepatic ER is present in males, as are the genes that encode for Vtg (Maitre, Mercier, Dole, & Valotaire, 1985; LeGuellec, Lawless, Valotaire, Kress, & Tinniswood, 1988). Consequently, the induction of Vtg in males and juvenile females has become a widely used and reliable biomarker for exposure to exogenous estrogens (Sumpter & Jobling, 1995; Folmar et al., 1996; 2001; Arukwe & Goksøyr, 2002; Jobling et al., 2002a; 2002b; Sole´ et al., 2002; ThomasJones et al., 2003). Hence, in fathead minnow males exposed to estrogens, plasma Vtg protein levels can attain the same range as in mature females (Korte et al., 2000; Kidd et al., 2007), though species differ in the magnitude of their vitellogenic response to exogenous estrogens (Walker et al., 1999; Tyler et al., 2005). The inducing actions on Vtg synthesis by mixtures of ERa agonists are additive (Thorpe et al., 2003; Brian et al., 2005), and even weak agonists can interact and contribute to the overall effectiveness of the mixture. Induction of Vtg is concentration-dependent until maximal levels are attained (Panter, Thompson, & Sumpter, 1998; Vajda et al., 2010). Maximal Vtg induction can occur through either intermittent or sustained exposure to exogenous estrogens (Panter, Thompson, & Sumpter, 2000). Depending upon the intensity of estrogenic exposure, maximal plasma Vtg concentrations may be reached after only three days of exposure (Lindholst, K. Pedersen, & S. Pedersen, 2000; Schultz, Orner, Merdink, & Skillman, 2001; Rose et al., 2002). Plasma Vtg in males declines upon depuration to clean water, although reduction to baseline concentrations may take many months because of slow Vtg clearance in males (Rodgers-Gray et al., 2001; Schultz et al., 2001; Hemmer et al., 2002). The specificity and time course of the vitellogenic response to exogenous estrogens in fishes have made it a useful biomarker of exposure to exogenous synthetic and biogenic estrogens. Elevated plasma Vtg has been detected in male fishes inhabiting estrogen-contaminated waters (Allen et al., 1999; Jobling et al., 1998; Van Aerle et al., 2001), in caged fishes exposed to WWTP effluents (Purdom et al., 1994; Harries et al., 1997), and in rainbow trout (Oncorhynchus mykiss) exposed under laboratory conditions to transported WWTP effluents (Harries et al., 1999) ¨ rn et al., and zebrafish exposed similarly to BKME (O 2006a). However, elevated Vtg in males is not in itself evidence of reproductive dysfunction (Mills et al., 2003). Although Vtg induction is an excellent marker of exposure to estrogenic compounds, links between protein induction and adverse effects at higher levels of biological organization, such as population impact, have not yet been

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established clearly (reviewed in Mills & Chichester, 2005). Elevated Vtg was not correlated with inhibition of malespecific sexual behavior in fathead minnows (Kramer, Miles-Richardson, Pierens, & Giesy, 1998) or sperm motility and fertilization capacity in male cunners (Tautogolabrus adspersus) (Mills et al., 2003). High Vtg levels in males could conceivably cause negative effects such as kidney dysfunction leading to death (Herman & Kincaid, 1988), and elevated plasma Vtg levels are correlated with increased mortality and decreased fitness in fathead minnows (Schmid, Gonzalez-Valero, Rufli, & Dietrich, 2002). Interpretation of elevated Vtg in free-living fishes is further complicated by the discovery that Vtg induction may follow exposure to a nonestrogenic precursor to estrogen biosynthesis, such as 17a-methyltestosterone (Ankley et al., 2002; Zerulla et al., 2002; Panter et al., 2004; Andersen et al., 2006) and, paradoxically, the nonaromatizable androgen dihydrotestosterone (Kim, Takemura, Kim, & Lee, 2003; Panter et al., 2004). Induction of Vtg by estrogens can be potentiated by cortisol (Brodeur, Woodburn, & Lecka, 2005). Induction of Vtg is antagonized by antiestrogens or androgens in medaka (O. latipes), sea bass (Dicentrarchus labrax), and zebrafish (Navas, ¨ rn, Yamani, & Norrgren, Zanuy, Segner, & Carrillo, 2004; O 2006), and reduced Vtg in female P. promelas has been used as a biomarker of exposure to environmental antiestrogens and androgens (Seki, Fujishima, Nozaka, Maeda, & Kobayashi, 2006; Villeneuve et al., 2009). Curiously, studies on exposures to BKMEs find a mixture of estrogenic effects such as induction of Vtg in immature female rainbow trout (Orrego et al., 2006), fathead minnows ¨ rn (Parrott, Wood, Boutot, & Dunn, 2003), and zebrafish (O et al., 2006a), although, most commonly, masculinization and/or depressed Vtg synthesis are observed in a number of ¨ rn et al., 2006b; Seki et al., 2006; see also review species (O by Hewitt et al., 2008). Some of these differences may relate to differences in the composition of the BKME from different mills; e.g., rainbow trout exposed to BKME in Chile (Oreggo et al., 2006) vs. rainbow trout exposed to a New Zealand mill effluent (Van den Heuvel, Ellis, ¨ rn et al. Tremblay, & Struthridge, 2002), although O (2006a) report both male-biased sex ratios and increased Vtg synthesis in zebrafish exposed to an effluent from a Swedish mill.

5.2.2. Central activational effects: feedback disruption The reproductive axis of vertebrates is regulated primarily via negative feedback by gonadal sex steroids. This is a mechanism exploited by pharmacological contraceptives in human females, where supraphysiological dosages of synthetic estrogens are administered to sexually mature


females to inhibit ovulation. In humans, the contraceptive function of EE2 is mediated via negative feedback by binding to hypothalamic ERs. Such interaction with negative-feedback circuitry inhibits pituitary release of the GTHs: FSH and LH. A similar dosing regimen occurs in fishes living downstream of human WWTPs, where treated wastewater effluents are discharged into surface waters (Larsson et al., 1999). Indeed, this extension of consequences of exposure from upstream to downstream organisms requires consideration of larger scale communication networks linking organisms of different species within a dynamic information network.

5.2.3. Activation of gametogenesis In teleost fishes, GTHs stimulate gonadal sex steroid biosynthesis and gametogenesis. Withdrawal of GTHs via negative feedback inhibits gametogenesis, reduces circulating sex-steroid concentrations, and reduces expression of steroid-dependent secondary sex characteristics. Disruption of gametogenesis in adult fathead minnows (MilesRichardson et al., 1999a; 1999b; Halm et al., 2002; Pawlowski, Van Aerle, Tyler, & Braunbeck, 2004; Leino, Jensen, & Ankley, 2005) and secondary sex characteristic expression (Miles-Richardson et al., 1999a; 1999b; Pawlowski et al., 2004; Parrott & Blunt, 2005) following exposure to EACs is likely an activational effect due to negative feedback and GTH withdrawal rather than a direct inhibitory action of EACs on peripheral tissues (Trudeau, 1997).

6. EVIDENCE OF REPRODUCTIVE DISRUPTION IN FREE-LIVING FISHES The western watersheds of North America are experiencing what is increasingly being recognized as the most severe and prolonged drought in 500 years (Cook, Woodhouse, Eakin, Meko, & Stahle, 2004). Low stream flows coupled with rapidly increasing human populations with increased water demands are reflected in the increases in number and size of communities discharging effluents into surface waters. Evidence of reproductive disruption is found downstream of the WWTPs of three different-sized Colorado communitiesdBoulder, Denver, and Colorado Springsdcompared to reference sites. These stream segments are close to the glacial and snowmelt headwaters of the Platte River and the Arkansas River. These communities represent the first major effluent discharges into these rivers, and during low-flow periods (most of the year) these downstream segments are characterized as effluent-dominated. Gonadal intersex, female-biased sex ratios, and other evidence of reproductive disruption consistent with exposure to exogenous estrogens occur in

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fishes below all three WWTPs sampled, but not at reference sites (Woodling et al., 2006; Vajda et al., 2008). Reproductive events of many mountain and plains river fishes are synchronized with stream flow patterns (Woodling, 1985). In headwater streams, spring snowmelt can lead to local flooding and dilution of effluent. However, low-flow conditions during late summer, fall, and winter lead to effluent domination of stream-flow below the WWTPs. Consequently, the highest concentrations of EAC contaminants in streams (Murphy et al., 2003) coincide with vulnerable periods of organizational development in many resident fish species (see review by Pandian & Sheela, 1995). Alterations to gonadal development and reproduction of wild fish populations around the world are associated with exposure to effluents originating from industrial (Larsson et al., 2000), and domestic sources (Sumpter & Jobling, 1995; Tyler & Routledge, 1998). The WWTP effluents of nonindustrial communities in consumer societies, which include naturally excreted steroids, pharmaceuticals, and personal care products, may be sufficiently endocrineactive to impact the reproduction of downstream organisms (Jobling et al., 2000a; 2002b; Sole´ et al., 2002; Vajda et al., 2008). These impacts on fish reproduction may occur in addition to more adverse consequences, such as overt toxicity and increased morbidity, for fishes downstream of WWTP effluents (Tsai, 1973; Fava & Tsai, 1976; Mitz & Giesy, 1985) or industrial discharges such as from paper and pulp mills (see Hewitt et al., 2008).

6.1. Waste Water Treatment Plant (WWTP) Effluent Interaction with Estrogen Signaling Networks Human populations, globally, discharge treated and untreated wastewater into surface waters, exposing downstream fishes and other organisms to synthetic and biogenic EACs. Most WWTPs target the reduction of pathogens, chlorine, phosphate, ammonia, and nitrogenous waste, but are not equipped for the specific reduction of synthetic or biogenic EACs. The majority of surface waters that receive treated municipal wastewater have detectable levels of steroidal and nonsteroidal estrogens (Johnson et al., 2005), androgens (Kirk, Tyler, Lye, & Sumpter, 2002; Thomas et al., 2002), and nonestrogenic neuroactive pharmaceuticals (Daughton & Ternes, 1999; Heberer, 2002), whereas upstream reference sites have much lower levels or undetectable levels of these EACs.

6.1.1. Steroidal estrogens in waste water treatment plant (WWTP) effluents Among the estrogens found in WWTP effluents are the naturally excreted steroidal estrogens E2, E1 and E3.

Even though WWTP processes are able to reduce the concentration of biogenic estrogens by 60e90% (reviewed in Christiansen, Winther-Nielsen, & Helweg, 2002), these compounds are found in WWTP effluents in biologically active (ng/L) concentrations (Desbrow, Routledge, Brighty, Sumpter, & Waldock, 1998; S. Snyder, Villeneuve, E. Snyder, & Giesy, 2001; Kolpin et al., 2002). Environmental persistence of steroidal estrogens is light- and temperature-dependent (Christiansen et al., 2002); instream microbial metabolism is able to convert most estrogens into nonestrogenic products (Christiansen et al., 2002).

6.1.2. Synthetic steroidal estrogens in waste water treatment plant (WWTP) effluents Waste water treatment plant effluents can contain the synthetic estrogen EE2 (Arcand-Hoy, Nimrod, & Benson, 1988; Larsson et al., 1999). 17a-ethinylestradiol is a synthetic steroidal estrogen used in most oral contraceptive pills. It is excreted in urine as free EE2 or as a glucuronideEE2 conjugate. This conjugate undergoes little degradation in municipal WWTP facilities, especially if sludge retention times are short (Andersen, Siegrist, Halling-Sorensen, & Ternes, 2003). Bacterial deconjugation of E2 and EE2 glucuronides during waste treatment release free E2 and EE2 to receiving waters (Adler et al., 2001; Andersen et al., 2003). Several examples of intersex formation following exposure to EE2 can be found in Table 13.1. Although E2 and EE2 are approximately equipotent in mammalian ER-based assays, EE2 appears to possess up to several orders of magnitude greater ER binding affinity and transcriptional activity in fishes (Rose et al., 2002; ThomasJones et al., 2003; Thorpe et al., 2003). The signaling challenge to aquatic biota posed by EE2 is enhanced by its environmental persistence; its environmental halflife is approximately 10 times that of E2 (17 days vs. 1.2 days, respectively) (reviewed in Christiansen et al., 2002).

6.1.3. Nonsteroidal estrogenic compounds in waste water treatment plant (WWTP) effluent Contributing to the endocrine activity of WWTP effluents are the nonsteroidal, estrogenic APs and alkylphenol ethoxylates (APEs). Alkylphenol ethoxylates are used as nonionic surfactants in diverse applications including soaps, detergents, and pesticide formulations. Alkylphenol ethoxylates are degraded to APs such as nonylphenol (NP) and octylphenol (OP). Although of lower estrogenic potency than steroidal estrogens (Purdom et al., 1994; Jobling, Reynolds, White, Parker, & Sumpter, 1995; Routledge et al., 1998), APs are discharged in WWTP effluents in substantially higher concentrations (Lye et al.,

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1999) and may contribute significantly to overall effluent estrogenicity (Jobling et al., 1998).

6.2. Masculinizing Effects of Endocrine-active Chemicals (EACs) Numerous studies have described masculinization of female fishes downstream of pulp and paper mill effluents in Europe, North America, and New Zealand (Davis & Bortone, 1992; Parks et al., 2001; Larrson, AdolphssonErici, & Thomas, 2006; Hewitt et al., 2008). Experimental assessments of the effects of BKME on female fishes indicate masculinization effects such as appearance of anal fin modification in the male direction in viviparous mosquitofish (Gambusia affinis) (Ellis et al., 2003), male-like coloration in guppies (Poecilia reticulata) (Larsson et al., 2002), nuptial tubercles in fathead minnows (Kovacs et al., 1995; Parrott et al., 2003), suppression of Vtg synthesis in longear sunfish (Lepomis megalotis) (Fentress, Steele, Bart, & Cheek, 2006), and male-biased sex ratios in eelpout (Zoarces viviparous) (Larsson et al., 2000) and zebrafish ¨ rn et al., 2006a). (O Trenbolone (TB) is a synthetic androgen that is implanted in feedlot steers to enhance growth. Cattle wastes find their way into nearby streams and rivers and represent a potential EAC. Treatment of adult female fathead minnows with low concentrations of TB results in development of male nuptial tubercles on the head (Ankley et al., 2003; Jensen, Makynen, Kahl, & Ankley, 2006; Seki et al., 2006; Martinovic et al., 2008a) and reduced Vtg secretion (Seki et al., 2006). Masculinization of female mosquito fish (Sone et al., 2005) and reduced Vtg secretion in medaka ¨ rn et al., 2006b) occurred as a result of TB and zebrafish (O exposure. Treatment of female medaka with TB or the P450aro inhibitor, fadrozole, reduced E2 levels and fecundity (Zhang et al., 2008). Several known antiandrogens have demasculinizing effects on fishes. Vinclozolin (VZ) is a fungicide that also has antiandrogenic actions. Although VZ reduced nuptial tubercles in male fathead minnows and blocked TB-induced nuptial tubercle formation fecundity in females, VZ paradoxically reduced fecundity in females and caused an upregulation of ARs (Martinovic et al., 2008a).

6.3. Pesticides in the Environment Agricultural pesticides reach the aquatic environment via runoff from rainfall and/or irrigation. One of the major pesticides of reproductive concern is atrazine, a triazine herbicide used primarily on corn and sorghum. Estrogenic effects of atrazine have been described in numerous amphibians (see Volume 2, Chapter 11), but no effects of triazine herbicides on reproduction were seen following short-term exposures of fathead minnows (Bringolf,


Belden, & Summerfelt, 2004; Villeneuve et al., 2006) or largemouth bass (Micropterus salmoides) (Martynuik, Sanchez, Szabo, Denslow, & Sepu´lveda, 2009). One 21-day study did report testicular disruption and ovarian atresia in sexually mature goldfish (Carassius auratus) treated with atrazine at 100 and 1000 parts per billion (ppb) (Spano et al., 2004), and the sensitivity to the female-priming pheromone, prostaglandin F2a, and the level of expressible sperm were reduced in mature Atlantic salmon parr (Salmo salar) by exposure to 1 or 2 ppb atrazine or the related triazine herbicide, simazine (Moore & Lower, 2001). Recent reviews of atrazine actions have concluded that it does not act as an estrogen agonist with respect to ERs (Eldridge et al., 2008) and that there is little support for it affecting reproduction of wild populations (Solomon et al., 2008). Nevertheless, atrazine appears to have other mechanisms capable of interfering with endocrine pathways by altering cyclic-3’,5’-adenosine monophosphate (cAMP) systems and disrupting reproduction (Suzawa & Ingraham, 2008). Atrazine exposure does impair salt balance and elevates plasma cortisol in juvenile Atlantic salmon (Waring & Moore, 2004; Nieves-Puigdoller, Bjo¨rnsson, & McCormick, 2007) and growth survival of the larval red drum (Sciaenops ocellatus) (McCarthy & Fuiman, 2008), suggesting other possible EAC roles for atrazine at environmentally relevant concentrations that might impair reproduction. Although the use of dichlorodiphenyltrichloroethane (DDT) was banned in the USA in 1972, DDT and its metabolites have persisted in the environment and still appear in surface and ground waters as well as in the bodies of vertebrates around the world (Hinck et al., 2007; Hinck et al., 2009; see also review by Guillette, Kools, Gunderson, & Bermudez, 2006; see also Volume 3, Chapter 14). Fishes captured in the ocean depths have been found to be contaminated with DDT (Storelli & Perrone, 2010). The most estrogenic form of DDT is the o,p0 -DDT isomer, and o,p0 -DDT is a more potent inducer of Vtg synthesis in rainbow trout hepatocytes than p,p0 -DDT (Okoumassoun et al., 2002). The major metabolites of DDT, p,p0 dichlorodiphenyldichloroethylene (DDE) and o,p0 -DDE, are estrogenic and antiandrogenic, respectively. Exposure to o,p0 -DDT (1e50 ppb) of 1-to-100-day-old medaka resulted in intersex production and enhanced Vtg production (Metcafe et al., 2000). Prolactin secretion by rainbow trout pituitaries is enhanced by o,p0 -DDT; this appears to work through ERs (Elango, Shepherd, & Chen, 2006). In contrast, a Florida population of the eastern mosquitofish (Gambusia holbrooki) shows no adverse reproductive effects of living a in a lake contaminated with DDT, DDE, and PCBs where alligators in the same lake exhibit considerable disruption (Edwards, Toft, & Guillette, 2010). Methoxychlor (MXC) became the replacement for DDT in part due to its being less persistent than DDT, although

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its chemical structure and biological actions are very similar. Although MXC is not estrogenic like DDT, once it enters the body of vertebrates, it is converted to metabolites that are estrogenic. Hence, we should refer to MXC as a proestrogen rather than as an estrogen. Treatment of male fish with MXC stimulates Vtg synthesis in carp (Cyprinus carpio) (Rankouhi et al., 2004), channel catfish (Ictalurus punctatus) (Nimrod & Benson, 1997), sheepshead minnow (Hemmer et al., 2001), and zebrafish (Ortiz-Zarragoitia & Cajaraville, 2005). In male largemouth bass (M. salmoides), MXC induced liver production of Vtg but inhibited ER expression in the testes (Blum, Nyagode, James, & Denslow, 2008). Secretion of T by minced ovaries from largemouth bass was also inhibited by MXC.

6.4. Neuroactive Pharmaceuticals in the Environment Many WWTP effluent-impacted surface waters also contain neuroactive psychotropic pharmaceuticals (Daughton & Ternes, 1999) that are capable of interacting with neural and neuroendocrine components of the fish reproductive axis (Mennigen et al., 2008). They accumulate in the brains of fishes downstream of WWTPs (e.g., Shultz et al., 2010) and can disrupt fish physiology and behavior at environmentally relevant concentrations (e.g., Painter et al., 2009). Serotonergic and catecholaminergic neurotransmitter systems regulate the responses of the fish HPG axis to physical (e.g., photoperiod and temperature) and biological (e.g., pheromones and visual signals) information, resulting in highly specific actions on reproductive development and behavior (Bla´zquez, Bosma, Fraser, Van Look, & Trudeau, 1998). In fishes, the synthesis of the neurotransmitters dopamine (DA), norepinephrine (NE), and serotonin (5-HT) is steroid-responsive and underlies the HPG feedback response to endogenous and exogenous estrogenic steroids (for review see Trudeau, 1997). This steroid responsiveness may contribute to the vulnerability of fish reproduction to disruption by neuroactive EACs such as fluoxetine and sertraline. Further, through direct interaction with brain neurotransmitter networks, environmental neuroactive chemicals can interfere with the sensing, integration, or transduction of biotic and abiotic information (Wingfield & Mukai, 2009).

6.5. Polycholorinated Biphenyls (PCBs) in the Environment The presence of PCBs in lake whitefish (Coregonus clupeaformis) (Mikaelian, De Fontaine, Harshbarger, Lee, & Martineau, 2002) and several introduced species of Pacific salmon (Leatherland, 1992) has been correlated with reproductive and thyroid abnormalities. Studies show that

PCBs are accumulated by fishes in part through their diet (Manenjian, O’Connor, Stewart, Miller, & Masnado, 2002), as well as by piscivorous consumers such as birds and mammals (e.g., Salata, Wade, Sericano, Davis, & Brooks, 1995; Bursian et al., 2006) including humans (Thomas & Colborn, 1992). Polychlorinated biphenyls can mimic thyroid hormones as well as produce direct and indirect estrogenic and antiestrogenic effects (McKinney & Waller, 1994; Connor et al., 1997; Kester et al., 2000). In mammals, including humans, PCBs obtained from consumption of Great Lakes fishes had significant detrimental effects on learning and memory (reviewed by Brouwer et al., 1999), but studies are needed to determine what neurological effects if any might occur in fishes. Interestingly, a recent study showing health benefits of fish consumption in the USA found that consumers of fishes from the Great Lakes Basin did not show these effects, emphasizing the deleterious effects of PCB and other EACs that are concentrated in fishes from this region (Tomasallo, Anderson, Haughwout, Imm, & Knobeloch, 2010).

7. CONCLUSIONS Overt evidence of endocrine disruption in fishes does not appear to be a ubiquitous and widespread environmental phenomenon, but is rather most likely to occur in populations exposed to endocrine-active source waters from WWTP effluents, pulp and paper mills, and populations in areas of high organic chemical contamination (Mills & Chichester, 2005). More widespread endocrine disruption may be observed in watersheds with smaller flows and correspondingly large or numerous endocrine-active inputs. Widespread endocrine disruption in fishes also may occur where habitats utilized by fishes during critical life stages are disproportionately impacted by endocrine-active effluents, even when overall watershed-scale endocrine impacts are low. The endocrine signaling networks that regulate reproduction in vertebrates have evolutionary roots that precede the origins of centralized neural tissues, endocrine organs, or vascular systems. Indeed, much of the subcellular and molecular infrastructure of vertebrate signaling networks has origins that precede multicellularity. Signaling by dietary and environmental chemicals likely played an important role in organismal signaling pathways prior to the origin of endogenous ligands (reviewed in Vajda & Norris, 2006). The openness of the vertebrate endocrine system to exogenous chemicals may represent an adaptation to permit the modulation of endocrine-mediated processes in response to diverse, meaningful environmental chemical signals. To the possible detriment of organismal fitness and global biological complexity, this openness extends to chemical misinformation or nonsense signaling, such as the signaling by EACs in anthropogenic contexts (reviewed in

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McLachlan, 2001). The consequences of disrupting organizational and activational reproductive processes may extend beyond the individual, with adverse effects emerging within social hierarchies, population growth, community structure, and ecosystem function.

ABBREVIATIONS 11-KT 5-HT AP APE AR BKME cAMP DA DDE DDT E1 E2 E3 EAC EAP EE2 ER FSH GnRH GTH HPA HPG HPI HPT LH MOA MXC NE NP OP P450aro PCB ppb T TB Vtg VZ WWTP

11-ketotestosterone Serotonin Alkylphenol Alkylphenol ethoxylate Androgen receptor Bleached kraft paper mill effluent Cyclic-3’,5’-adenosine monophosphate Dopamine Dichlorodiphenyldichloroethylene Dichlorodiphenyltrichloroethane Estrone 17b-estradiol Estriol Endocrine-active chemical Endocrine-active phytochemical 17a-ethinylestradiol Estrogen receptor Follicle-stimulating hormone Gonadotropin-releasing hormone Gonadotropin Hypothalamicepituitaryeadrenal Hypothalamicepituitaryegonadal Hypothalamicepituitaryeinterrenal Hypothalamicepituitaryethyroid Luteinizing hormone Mode of action Methoxychlor Norepinephrine Nonylphenol Octylphenol Aromatase Polychlorinated biphenyl Parts per billion Testosterone Trenbolone Vitellogenin Vinclozolin Waste water treatment plant

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Chapter | 13


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Species Index Index Abramus abrama, 251 Acanthopagrus schlegeli, 9, 34, 92, 156 Acipenser guldenstadti, 17 Acipenser ruthenus, 71 Acipenser stellatus, 91 Acipenser transmontanus, 18, 25 Aequidens pulcher, 131 Amblyraja radiata, 219 Amphiprion, 150, 151 Amphiprion melanopus, 155, 156 Amphiprion ocellaris, 10 Amphiprion percula, 65 Anabas testudineus, 92, 93 Anguilla anguilla, 20, 24, 66, 70, 93, 106, 161 Anguilla japonica, 34, 48 Apistogramma, 10 Apteronotus leptorhynchus, 133, 136 Astotilapia (Haplochromis) burtoni, 129, 131, 133, 157, 162, 179, 182, 183, 187 Astyanax, 186 Austichthys nobilis, 85 Barbonymus gonionotus, 135, 176 Barbus plebejus, 251 Bathygobius soporator, 129, 182 Beta splendens, 122, 129, 131 Brachyhypopomus, 136 Brachyhypopomus pinnicaudatus, 123, 124 Branchiostoma lanceolatum, 16 Calamoichthys calabaricus, 18 Callorhincus milii, 226, 230 Campylomormyrus, 124 Campylomormyrus compressirostris, 122 Campylomormyrus rhynchophorus, 122 Carassius auratus, 17, 70, 88, 107, 135, 169, 187, 254 Carassius carassius, 48, 170 Carcharius taurus, 210, 229 Carcharrhinus plumbeus, 211 Catostomus commersoni, 106, 251 Centrostephanus coronatus, 152 Cephalopholis boenak, 10 Cephaloscyllium laticeps, 219, 222 Channa gachua, 93 Cheilochromis euchilus, 187 Chelionichthys kumu, 110 Chlamydoselachus anguineus, 211 Chromis didpilus, 131 Cichlasoma bimaculatum, 135 Cichlasoma citrinellum, 10 Cichlasoma dimerus, 19 Clarias batrachys, 54, 135

Clarias gariepinus, 24, 50, 54, 55, 85, 162, 170, 251 Colisa lalia, 22, 133 Conger myriaster, 88 Coregonus clupeaformis, 19, 170, 177, 251, 255 Ctenopharyngodon idella, 177 Cyphotilapia, 183 Cyprinodon nevadensis amargosae, 133 Cyprinus carpio, 18, 89, 105, 170, 187, 255 Danio rerio, 19, 30, 44, 48, 66, 83, 105, 177, 250 Dasyatis akajei, 224 Dasyatus sabina, 212, 218, 222, 223, 224, 229, 230 Dicentrarchus labrax, 9, 19, 48, 66, 73, 85, 92, 111, 252 Diplodus puntazzo, 65 Eigenmannia, 136 Epalzeorhynchus frenatus, 177, 187 Epinephelus akaara, 92 Epinephelus coioides, 92 Epinephelus suillus, 154 Eptatratus burgeri, 194, 195, 196 Eptatratus deani, 194 Eptatratus stoutii, 194, 195, 196 Fugu ripes, 30 Fundulus, 66 Fundulus heteroclitus, 86, 134 Gadus morhua, 71 Gallus gallus, 30 Gambusia affinis, 254 Gasterosteus aculeatus, 4, 30, 49, 70, 122, 129, 130, 135, 251 Geotria australis, 93 Gibelion (Catla) catla, 52 Gillichthys mirabilis, 54 Gobius niger, 180 Gobiusculus flavescens, 125 Gramma loreto, 10 Gymnocephalus cernuus, 179 Haplochromis burtoni, 19 Hemichromis bimaculaus, 129 Hemichromis elongatus, 187 Hemigrammus, 186 Heterochromis multidens, 183 Heteropneustes fossilis, 26, 54, 55, 91 Hippocampus kuda, 47 Hippoglossus hippoglossus, 88

Hydrolagus colliei, 219 Hyphessobrycon, 186 Ictalurus nebulosus, 91 Ictalurus punctatus, 85, 90, 91, 154, 255 Jordanella floridae, 135 Labroides dimidiatus, 153 Labrus bergylta, 157 Lampetra aznandreai, 203 Lampetra fluviatilis, 202, 203 Lampetra japonica, 202 Latimeria, 240 Latimeria chalumnae, 45, 239 Latimeria menadoensis, 239 Latris lineate, 110 Lepidosiren paradoxa, 239, 241 Lepomis gibbosus, 129 Lepomis macrochirus, 51, 56 Lepomis megalotis, 129, 254 Lethrinops, 183 Lethrinops auritus, 187 Leucoraa ocellata, 228 Leucoraja (Raja) erinacea, 216, 217, 218, 219, 223, 230 Lobochilotes, 183 Lythripnis dalli, 152 Macropodus opercularis, 129, 135 Malacoraja senta, 219 Megalops atlanticus, 45 Megalops cyrpinoides, 170, 187 Menidia menidia, 9 Menidia peninsulae, 9 Micropogonias undulatus, 19, 49, 73, 162 Micropterus dolomieu, 251 Micropterus salmoides, 251, 254, 255 Misgurnus anguillicaudatus, 170, 176 Misgurnus fossilis, 135 Mordacia praecox, 198 Morone americana, 130, 134, 251 Morone saxatilis, 24, 85, 105 Mugil cephalus, 24, 72 Mugil planatus, 161 Mus musculus, 30 Mystus vittatus, 93 Myxine glutinosa, 194 Neoceratodus, 239, 240, 241, 242 Neoceratodus forsteri, 240, 241, 242, 243 Neogobius melanostomus, 54, 170, 179, 180, 187 Nerophis ophidion, 125 Nimbochromis polystugma, 187

265

266

Nonodelphis domestica, 30 Notopterus notopterus, 109 Odontesthes bonarensis, 10 Oncorhynchus, 66, 111, 178, 179 Oncorhynchus kisutch, 89, 93 Oncorhynchus masou, 18, 52, 93, 105 Oncorhynchus mykiss, 22, 48, 66, 83, 105, 130, 131, 133, 178, 251 Oncorhynchus nerka, 93, 111 Oncorhynchus nerka kennerlyi, 111 Oncorhynchus rhodurus, 85, 178 Oncorhynchus tshawytscha, 89, 111, 178 Opthalmotilapia nasuta, 183 Oreochromis, 24, 66 Oreochromis aureus, 182, 187 Oreochromis mossambicus, 121, 122, 154, 182 Oreochromis niloticus, 27, 68, 86, 133, 182 Oreochromis tanganicae, 182 Oreochromis upembae, 129 Orizias curvinotus, 2 Ornithorhynchus anatinus, 28 Oryzias latipes, 1, 2, 19, 30, 48, 66, 85, 109, 135, 252 Pagrus auratus, 107 Pagrus major, 24, 66, 70 Paralichthys olivaceus, 7, 10, 69, 88 Paramyzine atami, 196 Parapercis cylindrica, 110 Pelvicachromis, 10, 183 Pelvicachromis pulcher, 187 Peocilia phenops, 10 Perca fluviatilis, 251 Petromyzon marinus, 28, 66, 194, 197, 198 Pimephales promelas, 27, 48, 72, 88, 252 Platichthys flesus, 251 Platyrhinoidis triseriata, 212 Pleuronectes yokohamae, 251 Poecilia, 93 Poecilia reticulata, 93, 254 Polimyris isidori, 123 Pomacentris partitus, 162 Pomacentrus amboinensis, 108 Porites, 158 Proptopterus, 241, 242

Species Index

Proptopterus amphibious, 240 Proptopterus annectens, 240 Proptopterus dolloi, 240 Protopterus aethiopicus, 240 Pseudirasbora parva, 177 Pseudoanthias (Anthas) squampinnus, 151, 153 Pseudoblennius cottoides, 85 Pseudocrenilabrus, 183 Pseudocrenilabrus multicolor, 129, 131 Pseudotropheus zebra, 123 Psuedocrenilabrus multicolour, 76 Pundamilia, 183 Pundamilia nyererei, 187 Radulinopsis taranetzi, 121 Raja eglantera, 219 Rattus norvegicus, 30 Rhodeus oscellatus, 177 Rhodeus sericeus, 135 Rutilus rutilus, 177, 250, 251 Salaria pavo, 125, 126, 129, 132 Salmo salar, 48, 66, 88, 108, 131, 135, 170, 254 Salmo trutta, 106, 131, 170 Salvelinus, 178 Salvelinus alpinus, 55, 93, 107, 162, 170, 178 Salvelinus fontinalis, 178 Sarotherodon melanotheron, 129 Scaphirhynchus platorynchus, 251 Scartella cristata, 56 Schindleria praematura, 45 Sciaenops ocellatus, 254 Scophthalmus maximus, 88 Scyliorhinus canicula, 94, 216, 222, 226, 230 Silurus meridionalis, 8 Solea senegalensis, 27, 88, 162 Sparisoma viride, 154, 155, 156 Sparus aurata, 19, 24, 111 Sparus sarba, 105 Spathodus erythrodon, 183 Sphryna tiburo, 219, 223, 228, 230 Spondyliosoma cantharus, 121 Squalus, 220, 225 Squalus acanthias, 211, 212, 216, 217, 218, 219, 223, 224, 230

Steatocranus, 183 Stegastes leucostictus, 133, 134 Stegastes partitus, 123 Strongylocentrotus pururatus, 28 Sufflamen chrysopterum, 121 Sygnathus typhle, 125 Symphodus melops, 57, 121 Synganthus schlegeli, 47 Synodontis, 187 Takifugu niphobles, 1304 Takifugu rubripes, 4, 10, 21 Takydromus tachydromoides, 28 Tautogolabrus adspersus, 252 Tetraodon nigroviridis, 30 Thalassoma, 159, 160 Thalassoma bifasciatum, 10, 24, 127, 129, 150, 155, 156, 157, 158, 159, 160, 161, 164 Thalassoma duperrey, 56, 151, 154, 155, 157, 158, 159, 160, 161, 162, 163, 164 Thalassoma lucasanum, 159 Thunnus orientalis, 88 Thunnus thynnus, 48 Tilapia, 183 Tilapia zillii, 109, 121 Tinca tinca, 177 Torpedo, 220 Triakis semifasciata, 212 Trichogaster trichopterus, 129, 130 Trimma okinawae, 10, 152, 156, 157 Tropheus, 183 Tropheus moori, 187 Tylochromis sundanensis, 184, 187 Tyrranochromis nigroventris, 187 Urobatis (Urolophus) halleri, 212, 222 Xenopus, 74 Xenopus laevis, 28 Xenopus tropicalis, 28 Xenotilapia ornatipinnus, 183, 184, 187 Xiphophorus maculatus, 129 Zoarces viviparous, 251, 254 Zosterisessor ophiocephalus, 54

Subject Index Index ACTH, see Corticotropin Activin, oocyte maturation role, 75 Alkylphenol, estrogenic activity, 253 Alternative reproductive tactics (ARTs) overview, 126e127 testicular function, 56e57 AMH, see Anti-Mu¨llerian hormone g-Aminobutyric acid (GABA), gonadotropin release induction, 22e23 Androgen receptor (AR) brain expression, 33e34 endocrine-active chemicals, 248, 254 genomics studies, 49 Androstenedione lungfish function, 242 pheromone activity gobies, 181 males, 175e176 preovulatory steroid pheromone, 173e174 species specificity, 184 sex reversal role, 155 Anti-Mu¨llerian hormone (AMH), testicular differentiation role, 6e7 Appetitive/consummatory division, sexual behavior, 119e120 AR, see Androgen receptor Arginine vasotocin (AVT) chondrichthyan reproductive function, 225e226 sex reversal role, 157 sexual behavior role, 132e134 Aromatase brain expression, 31e32 male sexual behavior role, 128 oogonia differentiation role, 68e69 ovarian differentiation role, 7e9 sex reversal role, 156 temperature-dependent sex determination role, 9e10 ARTs, see Alternative reproductive tactics AVT, see Arginine vasotocin Calcitonin (CT), chondrichthyan reproductive function, 224 Cartilaginous fish, see Chondrichthyans Chondrichthyans arginine vasotocin in reproduction, 225e226 calcitonin in reproduction, 224 corticosterone in reproduction, 223 cycles of reproduction, 210e211 environmental effects on hormone levels, 229e230 female reproduction

steroid cycling and function dihydrotestosterone, 219 estradiol, 217e219 progesterone, 219 11-ketotestosterone, 219 steroidogenesis, 216e217 testosterone, 219 structure of reproductive tract, 215e216 hypothalamic-pituitary-gonadal axis gonadotropin-releasing hormone, 211e213 gonadotropins, 213e214 pituitary, 213e214 male reproduction testes structure and spermatogenesis, 219e220 steroid cycling and function dihydrotestosterone, 221e222 estradiol, 222e223 11-ketotestosterone, 221e222 progesterone, 223 testosterone, 221e222 steroidogenesis, 220e221 mating behavior, 211 prospects for study, 229e230 relaxin in reproduction, 223 reproductive behavior and sensory function, 229 reproductive modes, 209e210 serotonin in reproduction, 224e225 sex determination and maturation, 226e229 thyroid hormone in reproduction, 224 Coelacanths, see Sarcopterygians Corticosterone, chondrichthyan reproductive function, 223 Corticotropin (ACTH), functional overview, 104 Corticotropin-releasing factor (CRF), functional overview, 103e104 CRF, see Corticotropin-releasing factor CT, see Calcitonin CYP19, see Aromatase DDT, endocrine disruption, 254 Dehydroepiandrosterone sulfate (DHEAS), pheromone activity, 180 Deiodinases gonadal expression, 92e93 types and functions, 86e87 11-Deoxycorticosterone (DOC), gonadal effects, 107e108 DHEAS, see Dehydroepiandrosterone sulfate

DHT, see Dihydrotestosterone Dihydrotestosterone (DHT) brain metabolism, 34 chondrichthyan reproductive function females, 219 males, 221e222 dmrt1by sex determination, 2e4 testicular differentiation role, 5e6 dmy sex determination, 2e4 testicular differentiation role, 5 DOC, see 11-Deoxycorticosterone Dopamine functional diversity in teleosts, 24 inhibitory control of reproduction environmental cues in modulation, 26e27 evolutionary origins, 24 internal factors in modulation, 26 ovulation regulation, 76 pituitary gonadotropin control in teleosts, 23e24 sex reversal role, 157, 160e161 EACs, see Endocrine-active chemicals EGF, see Epidermal growth factor Electric organ discharge (EOD) mate selection role, 122e124 signal regulation, 136 Electro-olfactogram (EOG), pheromone response studies, 170e171, 178e184, 186, 188 Endocrine-active chemicals (EACs) activational disruption central feedback disruption, 252 gametogenesis activation, 252 vitellogenin induction in males, 251e252 androgenic compounds, 254 life stage-specific exposures, 248 multidimensional mixture complexity, 248 neuroactive pharmaceuticals, 255 organizational disruption gonadal differentiation, 249 gonadal intersex, 249e251 overview, 245e246 pesticides, 254e255 polychlorinated biphenyls, 255e256 signaling mechanisms androgen receptor, 248 estrogenic compounds estrogen additivity, 247e248 estrogen receptors, 247 steroidogenesis effects, 248

267

268

Endocrine-active chemicals (EACs) (Continued ) waste water treatment plant effluent nonsteroidal estrogens, 253e254 steroidal estrogens, 253 EOD, see Electric organ discharge EOG, see Electro-olfactogram Epidermal growth factor (EGF), oocyte maturation role, 75 ER, see Estrogen receptor Estradiol chondrichthyan reproductive function females, 217e219 males, 222e223 dopamine modulation in brain, 26 feedback loops, 33 female sexual behavior role, 132 glucuronide pheromones, 187 lampreys, 202e203 olfactory detection, 186 oogonia differentiation role, 68e69 ovarian differentiation role, 8 receptors, 26 spermatogenesis role, 49 stress effects, 110 thyroid function effects, 93 Estrogen receptor (ER) brain expression, 33 endocrine-active chemicals, 247e248 ovarian differentiation expression, 8 Estrone, pheromone activity, 180 Fertilization, internal versus external, 124e125 fMRI, see Functional magnetic resonance imaging Follicle-stimulating hormone (FSH) chondrichthyans, 213e214 hagfish, 196e197 lampreys, 199e200 oocyte maturation role, 74 oogenesis regulation, 70 spermatogenesis role, 44, 49 thyroid function effects, 94 Fox12, ovarian differentiation role, 8e9, 68e69 FSH, see Follicle-stimulating hormone Functional magnetic resonance imaging (fMRI), sexual behavior brain imaging, 137 GABA, see g-Aminobutyric acid GAL4-UAS, transgenic zebrafish, 138 Glycoprotein a2 (GPA2), functions, 16 Glycoprotein b5 (GPB5), functions, 16 Glycoprotein hormone (GpH), lamprey receptors, 200e202 GnIH, see Gonadotropin-inhibiting hormone GnRH, see Gonadotropin-releasing hormone Gonadal intersex, endocrine-active chemical induction, 249e251 Gonadotropin-inhibiting hormone (GnIH), regulators, 23 Gonadotropin-releasing hormone (GnRH)

Subject Index

brain distribution, 19e20 chondrichthyans, 211e213 discovery in teleosts, 18e19 genes, 19 hagfish, 196e197 lamprey functions, 198e199 receptor, 199 lungfish function, 242e243 pulsatile release studies, 21e22 receptors, 21 sequence comparison between species, 18 sex reversal role, 156, 161 sexual behavior role, 132e133 GPA2, see Glycoprotein a2 GPB5, see Glycoprotein b5 GpH, see Glycoprotein hormone GPR54 expression in puberty, 27 genes, 27 ligands, see Kisspeptins oocyte maturation role, 76 oogenesis role, 72 gsdf, testicular differentiation role, 5 Hagfish hypothalamic-pituitary-gonadal axis gonadotropins, 196e197 neurohypophysis and adenohypophysis, 196 ovary features, 195e196 overview of reproduction, 194e195 prospects for study, 204 secondary sexual characteristics, 196 sex determination, 195 testes features, 196 Hermaphroditism, see Gonadal intersex; Sex reversal Hydroxysteroid dehydrogenases brain expression and function, 34 stress effects, 110 11b-Hydroxytestosterone (OHT), male sexual behavior role, 128 Hypothalamic-pituitary-gonadal axis chondrichthyans gonadotropin-releasing hormone, 211e213 gonadotropins, 213e214 pituitary, 213e214 hagfish gonadotropins, 196e197 neurohypophysis and adenohypophysis, 196 lamprey glycoprotein hormone receptors, 200e202 gonadotropin-releasing hormone functions, 198e199 receptor, 199 gonadotropins, 199e200 neurohypophysis and adenohypophysis, 198 overview, 193e194

stress effects central nervous system, 105e106 gonadal function, 107e108 hepatic vitellogenesis effects, 106e107 pituitary function, 106 teleost features, 16e17 Hypothalamic-pituitary-intrarenal axis, see Stress response IGF-I, see Insulin-like growth factor-I Insulin-like growth factor-I (IGF-I) oocyte maturation role, 74e75 oogenesis regulation, 71 temperature-dependent sex determination role, 10 Isotocin (IST), sexual behavior role, 134e135 IST, see Isotocin 11-Ketosterone chondrichthyan reproductive function females, 219 males, 221e222 olfactory detection, 186 oogenesis regulation, 70e71 sex reversal role, 154e156 sexual behavior role female, 132 male, 128, 130e131, 137 spermatogenesis role, 49 stress effects, 107, 109e110 temperature-dependent sex determination role, 10 testicular differentiation role, 5 testicular synthesis, 47e48 Kisspeptins discovery in mammalian reproduction, 27 environmental regulation, 28 gene cloning, 28 link between growth/metabolism and reproduction, 30e31 oocyte maturation role, 76 origin and evolution, 28 receptor, see GPR54 sequence comparison between species, 29e30 teleost studies, 27e28 Lamprey circulating hormones during reproductive cycles, 203e204 gonadal development, 197e198 hypothalamic-pituitary-gonadal axis features glycoprotein hormone receptors, 200e202 gonadotropin-releasing hormone functions, 198e199 receptor, 199 gonadotropins, 199e200 neurohypophysis and adenohypophysis, 198 overview, 193e194 sex steroids, 202e203

269

Subject Index

life cycle, 197 ovary features, 198 prospects for study, 204 secondary sexual characteristics, 199 testes features, 198 LH, see Luteinizing hormone Lungfish, see Sarcopterygians Luteinizing hormone (LH) chondrichthyans, 213e214 hagfish, 196e197 lampreys, 199e200 oocyte maturation role, 74 oogenesis regulation, 70 postovulatory prostaglandin pheromone induction, 174e175 preovulatory steroid pheromone induction, 173e174 regulation of release, see also Gonadotropin-releasing hormone g-aminobutyric acid, 22e23 dopamine, 23e24 neuropeptide Y, 22 spermatogenesis role, 44, 49 stress effects, 109e110 thyroid function effects, 94 Mate selection, behavior, 122e124 Maturation-inducing hormone (MIH), oocyte maturation role, 73e74 Melatonin dopamine modulation in brain, 27 gonadotropin-inhibiting hormone regulation, 23 testicular function, 52e53 Methoxychlor endocrine disruption, 254e255 MIH, see Maturation-inducing hormone Neuropeptide Y (NPY), gonadotropin release induction, 22 Norepinephrine, sex reversal role, 160e163 NPY, see Neuropeptide Y OHT, see 11b-Hydroxytestosterone Olfaction pheromone detection, see Pheromones steroid glucuronide function, 55 Ovary differentiation, 6e9 hagfish features, 195e196 lamprey features, 198 oocyte maturation, 73e75 ovulation, 75e76 teleosts development stages, 66e68 germ cell differentiation into oogonia, 68e70 morphological aspects, 66 oogenesis regulation, 70e72 prospects for study, 76e77 reproductive strategies, 65e66 PACAP, see Pituitary adenylate cyclaseactivating polypeptide PCBs, see Polychlorinated biphenyls

PET, see Positron emission tomography Pheromones Atlantic salmon, 177e178 brown trout, 178 chars, 178 cichlids, 182e184 electro-olfactogram response, 170e171, 178e184, 186, 188 Eurasian ruffe, 179e180 gobies, 180e182 goldfish cypriniform pheromones in other than goldfish, 176e177 functional overview, 172e173 male pheromones, 175e176 postovulatory prostaglandin pheromone, 174e175 preovulatory steroid pheromone, 173e174 overview, 169e170 Pacific salmon, 178e179 phylogeny in salmonids, 179 species distribution, 171e172 species specificity, 184e188 Pituitary chondrichthyans, 213e214 hagfish, 196 lampreys, 198 teleosts, 17 Pituitary adenylate cyclase-activating polypeptide (PACAP), oocyte maturation role, 75 POA, see Preoptic area Polychlorinated biphenyls (PCBs), endocrine disruption, 255e256 Positron emission tomography (PET), sexual behavior brain imaging, 137 Postovulatory prostaglandin pheromone, 174e175 Preoptic area (POA) sex reversal role, 157, 164 sexual behavior role, 135e136 Preovulatory steroid pheromone, 173e174 Progesterone, chondrichthyan reproductive function females, 219 males, 223 Progestins male sexual behavior role, 131 pheromone activity cichlids, 182, 184 Eurasian ruffe, 180 preovulatory steroid pheromone, 173e174 structures, 187 Prostaglandins pheromones Atlantic salmon, 177 brown trout, 178 chars, 178 cypriniform pheromones, 176 Pacific salmon, 178

postovulatory prostaglandin pheromone, 174e175 sexual behavior role in teleosts, 135 Raphe nucleus (RN), sex reversal role, 163e164 Relaxin, chondrichthyan reproductive function, 223 Reproductive value (RV), sex reversal, 152e153 RN, see Raphe nucleus RV, see Reproductive value Saddleback wrasse, see Sex reversal SAH, see Size advantage hypothesis Sarcopterygians coelacanths, 240 lungfish aquaculture, 240e241 hormonal regulation of reproduction, 241e243 parental care, 241 spawning, 241 phylogenetic relationships, 239e240 prospects for study, 243 Seasonal reproduction, testicular function, 52e54 Seminal vesicles hormonal regulation, 54 plasma components and functions, 54e55 seasonal variation, 54 steroidogenesis, 54 Serotonin chondrichthyan reproductive function, 224e225 sex reversal role, 157e158, 160, 162e163 Sex determination chondrichthyans, 226e229 endocrine-active chemicals, 249 environmental effects on determination and differentiation, 9e10 environmental versus genetic, 1 genetics, 2e4 hagfish, 195 heterogametic modes, 2 prospects for study, 10e11 Sex reversal bidirectional sex reversal, 152 hermaphroditism in fish, 149e151 neuroendocrine regulation gonadal steroids, 154e156 monoamines, 157e158 peptides, 156e157 saddleback wrasse studies, 158e164 prospects for study, 164 protandrous sex reversal, 151e152 protogynous sex reversal, 151 role reversal overview, 125 social factors affecting, 153e154 theories, 152e153 SF-1, see Steroidogenic factor-1 Size advantage hypothesis (SAH), sex reversal, 152

270

Sox9 sex determination, 2, 4 testicular differentiation role, 6 Spawning lungfish, 241 site defense and preparation, 120e122 Spermatogenesis, see Testes SRY, sex determination, 2, 4 StAR, see Steroidogenic acute regulatory protein Steroidogenic acute regulatory protein (StAR), oogenesis regulation, 70e71 Steroidogenic factor-1 (SF-1), ovarian differentiation role, 9 Stress response effectors, 103e104 hypothalamic-pituitary-gonadal axis effects central nervous system, 105e106 gonadal function, 107e108 hepatic vitellogenesis effects, 106e107 pituitary function, 106 hypothalamic-pituitary-interrenal axis and reproductive response, 110e111 life stage-specific effects on reproduction adults, 109e110 larval stages, 108e109 puberty, 109 prospects for study, 112 reproduction and stress resistance, 111e112 Temperature, hormone response in chondrichthyans, 229e230 Temperature-dependent sex determination (TSD), mechanisms, 9e10 Territoriality, see Spawning Testes

Subject Index

accessory structures seminal vesicles, 53e55 testicular blind pouches, 53 testicular glands, 53 castration behavioral effects, 128e130 development, 51e52 differentiation, 4e6 genomics studies, 48e49 hagfish features, 196 hormone regulation of structure and function, 49e50 prospects for study, 57 seasonal aspects of function, 52e53 species differences in sperm characteristics and testicular function, 55e57 spermatogenesis, 44e45, 47, 49e50 sperm release timing, 52 steroid hormone synthesis, 47e48 types, 45e47 Testosterone chondrichthyan reproductive function females, 219 males, 221e222 female sexual behavior role, 131e132 lampreys, 202e203 lungfish function, 242 male sexual behavior role, 128e130 oogenesis regulation, 70e71 sex reversal role, 154e155 thyroid function effects, 93 TH, see Thyroid hormone Transthyretin (TTR), tissue distribution, 87 Thyroid hormone (TH) chondrichthyan reproductive function, 224 circulating level regulation, 84e87 clearance, 87 deiodinases, 86e87 functional overview, 83e84 receptors

T3, 88 T4, 88e89 reproductive function functional changes during reproductive maturation, 91e92 gonadal tissue regulatory elements, 92e93 manipulation studies, 94e95 overview, 90e91 sex steroids and thyroid function effects, 93e94 synthesis and release, 90 thyroid tissue features, 89e90 transporters, 87 types and structures, 83, 86 Thyroid-stimulating hormone (TSH) functional overview, 84e85 receptor, 85, 92 Thyrotropin-releasing hormone (TRH), functional overview, 84 Trembolone, androgenic activity, 254 TRH, see Thyrotropin-releasing hormone TSD, see Temperature-dependent sex determination TSH, see Thyroid-stimulating hormone TTR, see Transthyretin Vitellogenesis endocrine-active chemical induction in males, 251e252 estradiol role, 33 stress effects, 106e107 vitellogenin uptake, 71 Waste water treatment plant (WWTP), effluent neuroactive pharmaceuticals, 255 nonsteroidal estrogens, 253e254 steroidal estrogens, 253 WWTP, see Waste water treatment plant

Color Plates

FIGURE 1.1 Change in germ cell number and meiosisspecific marker gene expression in medaka XY embryos after dmy knock-down. dmy was knocked down using gripNAs in the XY embryos of medaka in order to ascertain its role during early stages of testicular development. dmy and human CREB (control) gripNAs were microinjected into single-cell-stage embryos, and the status of sexual differentiation in the gonads of these embryos was examined on the day of hatching by 80 histology and whole-mount in-situ hybridization XY(WMISH). XY and XY-hCREB fry had fewer germ cells, dmy while XX and XY-dmy possessed more germ cells, indicating initiation of ovarian differentiation in the gonads of the latter groups. Another indication of initiation of ovarian differentiation is the initiation of meiosis exclusively in 0 dah XX gonads. Here, the initiation of meiosis was assessed by WMISH with scp3 (photomi0 crographs). No signals could be detected for scp3 in XYXX XYXYXY hCREB dmy hCREB (upper panel), while strong expression of scp3 was observable in the gonadal region of XY-dmy (lower panel), confirming the entry of germ cells in these gonads into meiosis in the absence of Dmy. 160

Germ cell number

XYhCREB

XY

XX

FIGURE 1.2 Expression pattern of gsdf mRNA during early stages of sexual differentiation in medaka. Expression pattern of gsdf was examined in the XY and XX gonads on the day of hatching by in-situ hybridization using amplified gsdf aRNA probe. Predominant expression of gsdf was detectable in the somatic cells surrounding the germ cells in the XY gonads. Its expression was almost undetectable in the XX gonads of the same stage. Strong expression of gsdf only in the somatic cells of the presumptive testis during early sexual differentiation stages suggests a role for it in the initiation of Sertoli cell differentiation.

GSD

XY Initial trigger

FIGURE 1.6 Factors involved in genetic sex determination (GSD) and environmental sex determination (ESD) in teleosts (refer to the text for detailed description). 11-KT, 11-ketotestosterone; E2, 17b-estradiol.

ESD

XX Temperature pH Social factors

Dmy/Dmrt1bY Male pathway

Female pathway

Gsdf

Foxl2

Dmrt1

Ad4bp/sf-1

Sox9a2

Cyp19a1a

Male pathway

Female pathway

Dmrt1

Cyp19a1a

11-KT

E2

Major player Amh

Hormone

11-KT

E2

FIGURE 2.1 (a and b) Comparative organization of the hypothalamoepituitary connections in teleost fishes (a) and mammals (b). In fishes, there is no hypothalamusepituitary portal system. The hypophysiotropic neurons send their projections directly into the anterior lobe. Nerve endings either connect with secretory cells of the anterior lobe (1a; see (c) and (d)) or end at a basal membrane that separates the neurohypophysis (nh) from the adenohypophysis (1b). This innervation corresponds to that of the median eminence in mammals (1). In both fishes and mammals, magnocellular neurons (ocytocin/isotocin and vasopressin/vasotocin) send their projections to the neural lobe (¼ pars nervosa), which in teleost fishes is intermingled with the pars intermedia. (c and d) Examples of g-aminobutyric acid (GABA) terminals directly apposed to LHb (gonadotropin (GTH)) cells (a) or growth hormone (GH) cells (b). g-aminobutyric acid fibers were labeled by a preembedding technique using peroxidase, while pituitary hormones were labeled by postembedding immunohistochemistry with gold particles. Bar ¼ 0.5 mm. Reproduced from Kah et al. (1992), with permission from S. Karger AG, Basel.

FIGURE 2.2 (a) Hypothesis on the evolution of gonadotropin-releasing hormone (GnRH) genes in vertebrates based on the existence of a third full genome duplication, specific to teleost fishes (3R). This hypothesis postulates that invertebrates have at least one gnrh gene, leading to the expectation that the 1R and 2R duplications would have given birth to four potential gnrh genes, as is the case for GnRH receptor (GnRH-R) (see (b)). However, the current information suggests that one copy of these ancestral duplicated genes was rapidly lost. Thus, the current available information suggests that two gnrh genes, gnrh1 (green) and gnrh2 (purple), were present in basal representatives of both early actinopterygians and sarcopterygians. Following the teleost-specific genome duplication, a second copy of the gnrh1 gene emerged (gnrh3, red). However, in some teleost species, either gnrh1 or gnrh3 were lost. With respect to gnrh2, it is probable that the second copy gene was rapidly lost. (b) Hypothesis on the evolution of GnRH-Rs in vertebrates. Starting with one putative ancestral GnRH-R sequence in invertebrates, the 1R and 2R genome duplications potentially generated four sequences in basal sarcopterygians and actinopterygians. In support of this hypothesis, three GnRH-Rs were found in the bullfrog, a diploid species. In teleost fishes, the 3R teleost-specific genome duplication could have generated up to eight putative sequences; accordingly, five functional GnRH-R sequences have been demonstrated in certain teleost species. This indicates that other GnRH-R copies were lost along the way. Reproduced from Kah et al. (2007), with permission from Elsevier.

FIGURE 2.7 (a) Unrooted neighbor-joining bootstrap consensus tree of kiss1 and kiss2 genes in vertebrates. An alignment of amino acid sequences corresponding to exon 2 was used. Ambiguously aligned sequences and gaps were taken out before analysis. (b) Synteny of KiSS-1-containing regions in vertebrate genomes. In the mouse and rat, the proximity between KiSS-1 and GOLT1A (number 7) indicates that the sequence of KiSS-1 was found within GOLT1A. Genes shown in boxes in human chromosome 1 (Chr1) are located in other chromosomes in that species. (c) Synteny of KiSS-2-containing regions in vertebrate genomes. The genes boxed in the mouse and rat are located in the indicated chromosome. Dr, Danio rerio; Fr, Fugu rubripes; Ga, Gasterosteus aculeatus; Gg, Gallus gallus; Hs, Homo sapiens; Md, Monodelphis domestica; Mm, Mus musculus; Ol, Oryzias latipes; Rn, Rattus norvegicus; Tn, Tetraodon nigroviridis. In both schematic representations, numbers on the boxes indicate different genes. Orthologous genes are represented as squares, and those cases where paralogs were found in KiSS-1 and KiSS-2 analyzed regions are represented as ovals. A black bar between genes indicates that these are not contiguous. Dashed boxes for KiSS-1 or KiSS-2 represent absence of the gene. Numbers in brackets on the right indicate which chromosome (Ch), scaffold (S), or linkage group (LG) the cluster is found on. Reproduced from Felip et al. (2009), with permission from Elsevier.

FIGURE 4.1 General scheme of oocyte development and growth in fishes. The sequence of oocyte stages is as follows: stage Idprimary growth; stage IIdcortical alveoli growth period; stage IIIdearly vitellogenic oocytes; stage IVdlate vitellogenic phase; and stage Vdmature/ovulated oocyte, full of yolk (with lipid and protein globules). Oocyte growth is controlled by 17b-estradiol (E2) and follicle-stimulating hormone (FSH), whereas the resumption of meiosis is regulated by luteinizing hormone (LH) and maturation-inducing hormone (MIH). GVBD, germinal vesicle breakdown.

FIGURE 4.3 Overview of the physiological regulation of oocyte maturation in fishes. Luteinizing hormone (LH) is a key player during oocyte maturation that increases the activity of the maturation-inducing hormone receptor (MIH-R), via a still-unknown mechanism. Subsequently, MIH can bind to MIH-R and induce the maturation of oocytes. The LH signal is mediated by a local paracrine network that additionally triggers maturation. Insulin-like growth factor-I (IGF-I) increases the number of gap junctions and can independently induce the maturation of oocytes via the IGF-I receptor. The preovulatory surge of LH is triggered by positive feedback on GnRH in the brain. It is not known whether the KiSS-1 system is involved in this process in fishes as it is in mammals. 17a-OHP, 17a-hydroxyprogesterone; cAMP, cyclic-3’,5’-adenosine monophosphate; DA, dopamine; E2, 17b-estradiol; EGF, epidermal growth factor; GJ, gap junctions; GnRH, gonadotropin-releasing hormone; PKA, protein kinase A; PKC, protein kinase C; T, testosterone.

FIGURE 6.1 Overview of the major neuroendocrine signals and interactions between the hypothalamicepituitarye interrenal (HPI) stress axis (in red) and the hypothalamicepituitaryegonadal (HPG) reproductive axis (in blue) in teleosts. Neurotransmitters regulate the activity of both axes. Whereas activation of the HPI axis by stressors results in the production of cortisol by the interrenals, the HPG axis stimulates the production of the sex steroids, estradiol (E2), testosterone (T), and 11ketotestosterone (11-KT). Estradiol also stimulates the production of vitellogenin by the liver. Solid black arrows indicate stimulation. Dashed black arrows indicate inhibition. Red arrows indicate potentials effects of cortisol and corticotropin-releasing factor (CRF) on the HPG axis. Blue arrows indicate potential effects of sex steroids on the HPI axis. 5-HT, serotonin; ACTH, corticotropin; Ct, corticotropes; DA, dopamine; FSH, follicle-stimulating hormone; GABA, g-aminobutyric acid; GnRH, gonadotropinreleasing hormone; Gt, gonadotropes; LH, luteinizing hormone; NA, noradrenaline.

(a)

(b)

(c) FIGURE 7.4 Male alternative reproductive tactics in the peacock blenny Salaria pavo. Parasitic males (b) reproduce by mimicking the female (c) morphology and behavior in order to approach the nests of larger nesting males (a) and fertilize eggs.

FIGURE 10.2 Sower, Freamat, and Kavanaugh (2009) hypothesize that the hypothalamicepituitaryegonadal (HPG) and hypothalamice pituitaryethyroid (HPT) endocrine systems evolved from an ancestral, prevertebrate, exclusively neuroendocrine mechanism by gradual emergence of the components of a new control level (GpHs/GpH-Rs) concomitantly with the development of the corresponding anatomical structure (pituitary). The endocrine control of reproductive and thyroid functions in lampreys may reflect an intermediary stage on the evolutionary pathway to the highly specialized gnathostome HPG and HPT axes (Sower et al., 2009). CNS, central nervous system; GnRH, gonadotropin-releasing hormone; TRH, thyrotropin-releasing hormone; lGpH, lamprey glycoprotein hormone; lGpHR-I, lamprey glycoprotein hormone-receptor-I; lGpHR-II, lamprey glycoprotien hormone-receptor-II; T3, triiodothyronine; T4, thyroxine; CRH, corticotropin-releasing hormone; TSH, thyrotropin or thyroid stimulating hormone; GTH1, gonadotropin 1; GTH2, gonadotropin 2; FSH, follicle-stimulating hormone; LH, luteinizing hormone; FSH-R, follicle-stimulating hormone receptor; TSH-R, thyroid stimulating hormone receptor; LHR, luteinizing hormone receptor.

FIGURE 10.3 A proposed evolution of the glycoprotein hormone (GpH) subunits and glycoprotein hormone receptor (GpH-R) in vertebrates. (Bottom right) Freamat and Sower (2008) hypothesize that lGpH-R I and lGpH-R II are the only members of the GpH-R subfamily in lampreys. They are descendants of the thyrotropin receptor-like molecular ancestors of the GpH-Rs in gnathostomes and are likely the result of the genome duplication event hypothesized to have taken place before the divergence of lamprey lineage. (Bottom left) Ancestral glycoprotein subunits (GPA, a) and (GPB5, b) likely existed in a common ancestor of invertebrates and vertebrates (Sudo, Kuwabara, Park, Hsu, & Hsueh, 2005). Sower, Freamat, and Kavanaugh (2009) propose that an ancestral GpH gave rise to only one gonadotropin, and to the GpH family that gave rise to luteinizing hormone (LH), follicle-stimulating hormone (FSH), and thyroid-stimulating hormone (TSH) during the early evolution of gnathostomes. Lampreys may have one ancestral GPA2 and one, possibly two, beta subunits: GPb and/or GPB5 (unpublished data). The authors’ current working hypothesis is that the glycoprotein hormone subunit A2 identified is the ancestral a-subunit; after the gnathostomeeagnathan divergence, gene duplications produced the two a-subunits (GPA1 and GPA2) and three b-subunits (FSHb, LHb, and TSHb). Lampreys may have two GpHs: GPA2/b and/or GPA2/GPB5 (Sower et al., 2009). (Top) From the studies completed to date, Sower et al. (2009) hypothesize that there is lower specificity of gonadotropin (GTH) and its receptor in agnathans and that, during coevolution of the ligand and its receptor in gnathostomes, there were increased specificities of interactions between each GpH (TSH, LH, and FSH) and its receptor, depicted by the increase of shading from left to right. Thyrostimulin, another glycoprotein, is depicted by B5/A2 GpH. To date, thyrostimulin has only been shown to interact with TSH-R in gnathostomes. The question marks indicate the possible interactions of the subunits of the lamprey GpHs that are currently under investigation (Sower et al., 2009). LGR, Leucine-rich repeat-containing G-protein-coupled receptor.

FIGURE 12.2 Section of a mature ovary of the Australian lungfish (Neoceratodus forsteri) in the summer, undergoing regression. Portions of several large yolky oocytes are shown along with much smaller, unyolked, immature oocytes. The yolk is no longer evenly distributed throughout the regressing (atretic) oocytes. Many large macrophages are clustered alongside the oocytes, darkly pigmented following ingestion of yolk from unovulated follicles stained with Haematoxylin and eosin. Magnification 800.

Hormones and Reproduction of Vertebrates - Vol 1: Fishes

Hormones and Reproduction of Vertebrates - Vol 2: Amphibians

Hormones and Reproduction of Vertebrates - Vol 4: Birds

Hormones and Reproduction of Vertebrates - Volume 4: Birds

Hormones and Reproduction of Vertebrates - Volume 5: Mammals

Hormones and Reproduction of Vertebrates - Volume 3: Reptiles

Compendium of Trace Metals and Marine Biota: Vol 1: Plants and Invertebrates Vol 2: Vertebrates

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Hormones and Their Actions: Specific Actions of Protein Hormones

Hormones and Signaling

Nematode Parasites of Vertebrates

Growth and Lactogenic Hormones

Vertebrados - Vertebrates

Primitive Fishes (Fish Physiology, Vol. 26)

Chemical ecology of vertebrates

Loaves and Fishes

Best of Vol 1

Hormones and brain plasticity

Hormones And Their Receptors In Fish Reproduction (Molecular Aspects of Fish & Marine Biology)

Infertility and Assisted Reproduction

Interrelationships of Fishes

Migration of Freshwater Fishes

Migration of Freshwater Fishes

The Toxicology of Fishes

Molecular Systematics of Fishes

Interrelationships of Fishes

Freshwater Fishes of Iraq

Shore Fishes of Hawai'i

Molecular Systematics of Fishes

Hormones and Reproduction of Vertebrates - Vol 1: Fishes

Hormones and Reproduction of Vertebrates - Vol 2: Amphibians

Hormones and Reproduction of Vertebrates - Vol 4: Birds

Hormones and Reproduction of Vertebrates - Volume 4: Birds

Hormones and Reproduction of Vertebrates - Volume 5: Mammals

Hormones and Reproduction of Vertebrates - Volume 3: Reptiles

Compendium of Trace Metals and Marine Biota: Vol 1: Plants and Invertebrates Vol 2: Vertebrates

Vertebrates

Fishes

Hormones and Their Actions: Specific Actions of Protein Hormones

Hormones and Signaling

Nematode Parasites of Vertebrates

Growth and Lactogenic Hormones

Vertebrados - Vertebrates

Primitive Fishes (Fish Physiology, Vol. 26)

Chemical ecology of vertebrates

Loaves and Fishes

Best of Vol 1

Hormones and brain plasticity

Hormones And Their Receptors In Fish Reproduction (Molecular Aspects of Fish & Marine Biology)

Infertility and Assisted Reproduction

Interrelationships of Fishes

Migration of Freshwater Fishes

Migration of Freshwater Fishes

The Toxicology of Fishes

Molecular Systematics of Fishes

Interrelationships of Fishes

Freshwater Fishes of Iraq

Shore Fishes of Hawai'i

Molecular Systematics of Fishes

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