Nervous Systems and Control of Behavior (The Natural History of the Crustacea) [3, 1 ed.] 0199765677, 9780199765676, 0199791716

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Nervous Systems and Control of Behavior

The Natural History of the Crustacea Series Series Editor: Martin Thiel Editorial Advisory Board: Geoff Boxshall, Natural History Museum, London, UK Emmett Duffy, Virginia Institute of Marine Sciences, Gloucester, USA Darryl Felder, University of Louisiana, Lafayette, USA Gary Poore, Victoria Museum, Melbourne, Australia Bernard Sainte-Marie, Fisheries and Oceans Canada, Mont-Joli, Canada Gerhard Scholtz, Humboldt University Berlin, Berlin, Germany Fred Schram, Friday Harbor Marine Laboratory, Seattle, USA Les Watling, University of Hawaii, Hawaii, USA Functional Morphology and Diversity (Volume 1) Edited by Les Watling and Martin Thiel Lifestyles and Feeding Biology (Volume 2) Edited by Martin Thiel and Les Watling Nervous Systems and Control of Behavior (Volume 3) Edited by Charles Derby and Martin Thiel

Nervous Systems and Control of Behavior The Natural History of the Crustacea, Volume 3

EDITED BY CHARLES DERBY AND MARTIN THIEL

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1 Oxford University Press is a department of the University of Oxford. It furthers the University’s objective of excellence in research, scholarship, and education by publishing worldwide. Oxford New York Auckland  Cape Town  Dar es Salaam  Hong Kong  Karachi Kuala Lumpur Madrid Melbourne Mexico City Nairobi New Delhi Shanghai Taipei Toronto With offices in Argentina Austria Brazil Chile Czech Republic France Greece Guatemala Hungary Italy Japan Poland Portugal Singapore South Korea Switzerland Thailand Turkey Ukraine Vietnam Oxford is a registered trademark of Oxford University Press in the UK and certain other countries. Published in the United States of America by Oxford University Press 198 Madison Avenue, New York, NY 10016 © Oxford University Press 2014 All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, without the prior permission in writing of Oxford University Press, or as expressly permitted by law, by license, or under terms agreed with the appropriate reproduction rights organization. Inquiries concerning reproduction outside the scope of the above should be sent to the Rights Department, Oxford University Press, at the address above. You must not circulate this work in any other form and you must impose this same condition on any acquirer. Library of Congress Cataloging-in-Publication Data Nervous Systems and Control of Behavior / edited by Charles Derby and Martin Thiel. pages cm.—(The natural history of the crustacea ; volume 3) Includes bibliographical references and index. ISBN 978–0–19–979171–2 (alk. paper) 1.  Crustacea—Nervous system.  I.  Derby, Charles (Charles Dorsett), editor of compilation.  II.  Thiel, Martin, 1962– editor of compilation. QL935.C78 2014 595.315—dc23 2014014228

9 8 7 6 5 4 3 2 1 Printed in the United States of America on acid-free paper

PREFACE

Our collaboration on this project had its origin at a quite fitting location—The Crustacean Society Summer Meeting at the Tokyo University of Marine Science and Technology in Japan, in September 2009. Amid the symposia and poster presentations, we discussed the possibility of working together on this project. Later that winter, in Atlanta, we completed our initial plans and outline for the book. Our joint enthusiasm for crustaceans, enriched by our different perspectives and approaches, has fueled our perseverance toward the project since then. The outcome is this book—Nervous Systems and Control of Behavior. It is the third volume of a ten-volume series titled The Natural History of the Crustacea. Our volume builds on the foundation of the first two, which deal with more general topics: Volume 1: Functional Morphology and Diversity, and Volume 2: Life Styles and Feeding Biology, both edited by Les Watling and Martin Thiel. The next seven volumes will focus on various aspects of the lives of crustaceans, including growth, reproduction, development, life history and behavioral ecology, evolution and biogeography, fisheries and aquaculture, and ecology and conservation biology. We look forward to the completion of series, and we hope that the academic community finds the series useful.

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ACKNOWLEDGMENTS

We thank our contributors for sharing their knowledge and time in preparing their chapters. All are leaders in their respective disciplines, and thus have very busy professional lives, so finding the time for a project such as this one is challenging. We appreciate their willingness to join in our shared endeavor. Special thanks to our editorial assistants, Lucas Eastman and Annie Mejaes, who provided outstanding help in organizing, managing, and editing. The generous contribution from Universidad Católica del Norte was essential for this project—we are grateful for the continuous support that allowed us to focus on the task. The vision and foresight of the university authorities made this project possible, and we hope that this and the upcoming volumes will fulfill their expectations. We thank our external referees, who provided valuable comments to our authors and us about their chapters. We recognize our publisher, Oxford University Press, for its commitment to the project. We want to recognize all of those scientists who have contributed over the many years to our knowledge of crustacean neurobiology. Some of their work is referenced in this volume, but unfortunately many references to outstanding contributions are uncited here because of page restrictions (as our contributors well know!). Finally, we thank our lab members past and present for their ideas, energy, activities, and camaraderie, and our families for their support, encouragement, and endurance. Editing of this book was generously supported by Universidad Católica del Norte, Chile

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CONTRIBUTORS

EDITORS Charles D. Derby Neuroscience Institute and Department of Biology Georgia State University P.O. Box 5030 Atlanta, GA 30302 USA Martin Thiel Facultad Ciencias del Mar Universidad Católica del Norte Larrondo 1281 Coquimbo Chile AUTHORS Harold Atwood Department of Physiology Medical Sciences Building University of Toronto 1 King’s College Circle Toronto, Ontario M5S 1A8 Canada Ted Brookings Volen Center, Mailstop 013 Brandeis University 415 South Street Waltham, MA 02454 USA

Jonathan Caplan Volen Center, Mailstop 013 Brandeis University 415 South Street Waltham, MA 02454 USA Daniel Cattaert Institut de Neurosciences Cognitives et Intégratives d’Aquitaine CNRS & Université de Bordeaux PAC Talence Allée Geoffroy Saint-Hilaire Bât. B2 4ème étage Est CS 50023 33615 Pessac Cedex France Thomas W. Cronin Department of Biological Sciences University of Maryland, Baltimore County 1000 Hilltop Circle Baltimore, MD 21250 USA Charles D. Derby Neuroscience Institute and Department of Biology Georgia State University P.O. Box 5030 Atlanta, GA 30302 USA

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x Contributors Donald H. Edwards Neuroscience Institute Georgia State University 850 Petit Science Center Atlanta, GA 30303 USA David A. Ernst Department of Biology University of North Carolina Coker Hall CB-3280 Chapel Hill, NC 27599 USA María Luisa Fanjul-Moles Departamento de Ecología y Recursos Naturales Laboratorio de Neurofisiología Comparada Facultad de Ciencias, Universidad Nacional Autónoma de México Avenida Universidad 3000 Ciudad Universitaria México City 04510 México Kathryn D. Feller Department of Biological Sciences University of Maryland, Baltimore County 1000 Hilltop Circle Baltimore, MD 21250 USA Raymon Glantz Department of Biochemistry and Cell Biology Rice University Houston, TX 77251 USA Marie L. Goeritz Volen Center, Mailstop 013 Brandeis University 415 South Street Waltham, MA 02454 USA Gabrielle J. Gutierrez Group for Neural Theory Département d’Etudes Cognitives Ecole Normale Supérieure

29 rue d’Ulm 75005 Paris France Sara Haddad Volen Center, Mailstop 013 Brandeis University 415 South Street Waltham, MA 02454 USA Albert Hamood Volen Center, Mailstop 013 Brandeis University 415 South Street Waltham, MA 02454 USA Ronald Harris-Warrick Department of Neurobiology and Behavior Cornell University Ithaca, NY 14853 USA Daniel K. Hartline Békésy Laboratory of Neurobiology Pacific Biosciences Research Center University of Hawai’i at Manoa 1993 East-West Road Honolulu, HI 96822 USA Steffen Harzsch Zoologisches Institut und Museum Cytologie und Evolutionsbiologie Universität Greifswald Soldmannstrasse 23 D - 17487 Greifswald Germany William J. Heitler School of Biology University of St Andrews Fife KY16 9TS UK Jan M. Hemmi School of Animal Biology and the UWA Oceans Institute

Contributors University of Western Australia 35 Stirling Highway Crawley, WA 6009 Australia Jens Herberholz Department of Psychology Neuroscience and Cognitive Science Program University of Maryland 2123H Biology-Psychology Building College Park, MD 20742 USA Ronald R. Hoy Cornell University Department of Neurobiology and Behavior S.G. Mudd Hall Ithaca, NY 14853 USA Bruce R. Johnson Cornell University Department of Neurobiology and Behavior S.G. Mudd Hall Ithaca, NY 14853 USA Matthes Kenning Cytologie und Evolutionsbiologie Zoologisches Institut und Museum Soldmannstrasse 23 D - 17487 Greifswald Germany Tilman Kispersky Volen Center, Mailstop 013 Brandeis University 415 South Street Waltham, MA 02454 USA Franklin B. Krasne Department of Psychology and Brain Research Institute University of California, Los Angeles 1285 Franz Hall

Los Angeles, CA USA Petra H. Lenz Békésy Laboratory of Neurobiology Pacific Biosciences Research Center University of Hawaii at Manoa 1993 East-West Road Honolulu, HI 96822 USA Gregory Lnenicka Department of Biological Sciences University at Albany, State University of New York 1400 Washington Avenue Albany, NY 12222 USA Kenneth J. Lohmann Department of Biology University of North Carolina Coker Hall CB-3280 Chapel Hill, NC 27599 USA Héctor Maldonado (deceased), but formerly at: Universidad de Buenos Aires Argentina Eve Marder Volen Center, Mailstop 013 Brandeis University 415 South Street Waltham, MA 02454 USA DeForest Mellon, Jr. University of Virginia Department of Biology 288 Gilmer Hall 485 McCormick Road Charlottesville, VA 22903 USA David C. Sandeman Cytologie und Evolutionsbiologie Zoologisches Institut und Museum

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xii Contributors Soldmannstrasse 23 D - 17487 Greifswald Germany Manfred Schmidt Neuroscience Institute and Department of Biology Georgia State University P.O. Box 5030 Atlanta, GA 30302 USA Sonal Shruti Volen Center, Mailstop 013 Brandeis University 415 South Street Waltham, MA 02454 USA

IFIBYNE-CONICET Pabellón 2 Ciudad Universitaria (1428) Intendente Güiraldes 2160 Ciudad de Buenos Aires Argentina Marc J. Weissburg School of Biology Georgia Institute of Technology 310 Ferst Drive Atlanta, Georgia 30332 USA Robert A. Wyttenbach Neuroscience and Behavioral Biology Emory University 1462 Clifton Road NE Atlanta, GA 30322 USA

Martin Thiel Facultad Ciencias del Mar Universidad Católicadel Norte Larrondo 1281 Coquimbo Chile

Cornell University Department of Neurobiology and Behavior S.G. Mudd Hall Ithaca, NY 14853 USA

Daniel Tomsic Laboratorio de Neurobiología de la Memoria, Departamento Fisiología, Biología Molecular y Celular Facultad de Ciencias Exactas y Naturales Universidad de Buenos Aires

Jochen Zeil Research School of Biology The Australian National University Building 46, Biology Place Canberra, ACT 0200 Australia

CONTENTS

  1.  Nervous Systems and Control of Behavior: An Introduction   •  1 Charles D. Derby and Martin Thiel    2.  Adaptive Trends in Malacostracan Brain Form and Function Related to Behavior   •  11 David C. Sandeman, Matthes Kenning, and Steffen Harzsch    3.  Sensory Systems of Crustaceans   •  49 DeForest Mellon, Jr.    4.  Peripheral Components of Crustacean Motor Systems   •  85 Harold Atwood    5.  Modulation of Crustacean Networks for Behavior   •  114 Ronald Harris-Warrick    6.  Synapses in Crustaceans   •  147 Gregory Lnenicka    7.  Adult Neurogenesis in Crustaceans   •  175 Manfred Schmidt    8.  Visual Systems of Crustaceans   •  206 Raymon Glantz    9.  Sensory Ecology of Vision in Crustaceans   •  235 Thomas W. Cronin and Kathryn D. Feller 10.  The Chemical Senses and Chemosensory Ecology of Crustaceans   •  263 Charles D. Derby and Marc J. Weissburg

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xiv Contents 11.  Mechanoreception in Crustaceans of the Pelagic Realm   •  293 Petra H. Lenz and Daniel K. Hartline 12 The Geomagnetic Sense of Crustaceans and Its Use in Orientation and Navigation   •  321 Kenneth J. Lohmann and David A. Ernst 13.  The Crustacean Stomatogastric Nervous System   •  337 Eve Marder, Marie L. Goeritz, Gabrielle J. Gutierrez, Albert Hamood, Ted Brookings, Jonathan Caplan, Sara Haddad, Tilman Kispersky, and Sonal Shruti 14.  Neural Control of Posture and Walking in Crustaceans   •  362 Daniel Cattaert 15.  The Escape Behavior of Crayfish   •  396 Franklin B. Krasne, William J. Heitler, and Donald H. Edwards 16.  Biological Rhythms and Their Neural Basis in Crustaceans   •  428 María Luisa Fanjul-Moles 17.  Neurobiology of Social Status in Crustaceans   •  457 Jens Herberholz 18.  Path Integration, Vision, and Decision-Making in Fiddler Crabs   •  484 Jochen Zeil and Jan M. Hemmi 19.  Neurobiology of Learning and Memory of Crustaceans   •  509 Daniel Tomsic and Héctor Maldonado 20. Crustaceans as Model Systems for Teaching Neuroscience: Past, Present, and Future   •  535 Bruce R. Johnson, Robert A. Wyttenbach, and Ronald R. Hoy Index   •  555

Nervous Systems and Control of Behavior

1 NERVOUS SYSTEMS AND CONTROL OF BEHAVIOR: AN INTRODUCTION

Charles D. Derby and Martin Thiel

Abstract Crustaceans have been favored by neuroscientists as experimental animals due to their diversity, interesting and complex behaviors, and accessible nervous systems. Since the 19th century, researchers have used crustaceans to understand fundamental properties of neurons, neural networks, chemical transmission and modulation, and other features of nervous systems, including insight into our own nervous systems in health and disease. Other neuroscientists have focused more on understanding the natural behavior of crustaceans and the underlying neural mechanisms. This chapter introduces topics covered under these two major themes and as presented in the chapters of our volume, including a historical context, and looks forward by considering future opportunities and challenges for crustacean neuroscience.

HISTORY OF CRUSTACEAN NEUROSCIENCE Crustaceans have been used as model organisms in all fields of biology, including neurobiology, developmental biology, physiology, evolutionary ecology, biogeography, and resource management. One reason for this is the huge number of crustacean clades and species and their incredibly disparate and diverse forms (sensu Schram 2012), arguably the greatest of any group of animals. With approaching 70,000 described extant species and with estimates of the actual number being many times this (Martin and Davis 2007, Ahyong et al. 2011), they range in size from microscopic (less than one mm in length and weighing less than one gram) to enormous (nearly one meter in length and weighing over 20 kg). Their disparity and diversity is reflected by their adaptations to the various environments that they inhabit, from dry deserts to deepsea hydrothermal vents. This disparity and diversity allows exploration of many issues, from the

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Charles D. Derby and Martin Thiel comparative to the biomedical to the applied. The fact that some species have large individuals that are commercially important for their edibility allows these species to be available in larger numbers, which is essential for some types of research. Figure 1.1 depicts the phylogenetic diversity of crustaceans and their affiliation with other arthropods such as hexapods (insects) and chelicerates, which have also been used extensively in neurobiological research (e.g., see Strausfeld 2012). The vast majority of neurobiological research on crustaceans has been performed on the Malacostraca and secondarily on the Maxillopoda, especially the Copepoda. Within the Malacostraca (center panel of Fig. 1.1), most neurobiological work has been on the Decapoda, and secondarily on Mysidacea, Amphipoda, Isopoda, and Stomatopoda. Within the Decapoda (right panel of Fig. 1.1), most neurobiological research has been conducted on the Brachyura (true crabs), Achelata (especially spiny lobsters), Homarida (clawed lobsters), Astacida (crayfish), and Anomala (hermit crabs and squat lobsters). Neuroscientific research on crustaceans has a long history, some of it done by famous scientists. Gustaf Retzius of Sweden (for whom the huge Retzius cell of leeches and Cajal-Retzius cell in embryonic humans and other mammals are named) published in 1890 the first neuroanatomical description of the central nervous system of crayfish with neuronal resolution. Sigmund Freud, while still a student at the University of Vienna, studied the neuronal cytoskeleton using cells in the crayfish ventral nerve cord. He described the cellular organization of microtubules, though they were not called that at the time, in a publication of 1882. Albrecht Bethe of Germany (who is perhaps better known as the father of Hans Albrecht Bethe, the Nobel laureate nuclear physicist) published a series of four papers in the 1890s on the neuroethology of the shore crab Carcinus maenas (1895, 1897a, 1897b, 1898). He attempted to produce a complete neuroanatomical description of this crab’s central nervous system and, based on that, he performed experiments in an attempt to explain the neural basis of several behaviors. As techniques, he used neuroanatomical staining, primarily with methylene blue, and microsurgical ablations to study movement of appendages, principally the antennae and eyestalks, but also walking legs. He succeeded in identifying motor nerves underlying these movements, and to some degree the location of the motor neuronal somata and their neuropil arborizations. He not only performed microsurgical ablations but also evaluated the success of his ablations through microscopical analysis of serial sections through the treated brains. He had several interesting findings. One was that the motor neuronal somata are not necessary for the functioning of these cells. Another is that in male crabs, the olfactory lobes control discrimination between postmolt females and other animals or objects. Normal crabs only copulated with postmolt females; crabs without olfactory lobes copulated with every crab or object of about the right size. Based on results such as these, Bethe concluded that the brain is a “reflex-inhibiting” organ (he called these behaviors “reflexes”), and that the olfactory lobes are primarily responsible for this activity. Without the correct connections in the brain, reflexive behaviors such as feeding, copulation, or grooming do not stop at the appropriate time and thus become maladaptive. By the end of his studies, Bethe realized that neither the staining technology available to him nor the resolution of his ablation experiments matched the complexity of the central nervous system and the behaviors produced by it. Consequently, despite his choice to work on a supposedly “simple” organism, he became frustrated and concluded his work with the following statement: “So bin ich auch jetzt, wo ich meine ganze Arbeit überblicke, zu dem traurigen Befunde gekommen, dass für die faktische Erkenntnis nichts dadurch gewonnen ist. Fände man nicht Befriedigung in dem Suchen nach Erkenntnis, man würde die Hände verzweifelnd in den Schoss legen und sagen: Es ist zu schwer für uns Menschen” (Bethe 1898). [Translation: Now that I look back on the entirety of my work, I  come to the sad realization that the factual knowledge has not resulted in major conclusions. If not for the satisfaction in the search for knowledge, I might throw my hands up in desperation and say, “It is too difficult for us humans.”] Despite this expression of frustration,

Fig. 1.1. Phylogenetic diversity of crustaceans. This phylogenetic representation is a composite of those of Regier et  al. (2010) (left panel) and Scholtz and Richter (1995), Richter and Scholtz (2001), and Wirkner and Richter (2010) (center and right panels). There are alternative interpretations (e.g., Martin and Davis 2001, Porter et al. 2005, Tsang et al. 2008, Bracken et al. 2009, von Reumont et al. 2012) with minor deviations in particular clades, but the general relationships shown in the figure reflect most current phylogenies. insects

Hexapoda

cephalocarids, remipedes

Xenocarida

Malacostraca

copepods, cirripedes (barnacles)

Maxillopoda

anostracans (fairy shrimp, brine shrimp), cladocerans (water fleas), notostracans (tadpole shrimp)

Branchiopoda

ostracods, mystacocarids, brachiurans

Oligostraca

centipedes, millipedes

Myriapoda

horseshoe crabs, spiders, scorpions, mites, sea spiders

Chelicerata+Pycnogonida

Isopoda

Cumacea

Amphipoda

Mysidacea+Lophogastrida

thermosbaenaceans

Pancarida

Euphausiacea

anaspidaceans

Syncarida

Decapoda

mantis shrimp

Stomatopoda

phyllocarids

Leptostraca

Reptantia

Eumalacostraca

Pancrustacea Crustacea

Mandibulata

Arthropoda

true crabs

Brachyura

hermit crabs, squat lobsters, coconut crab, mole crabs

Anomala

mud shrimp, ghost shrimp

Thalassinida

crayfish

Astacida

clawed lobsters

Homarida

spiny lobsters, slipper lobsters

Achelata

Polychelida

Stenopodidea

caridean shrimp

Caridea

shrimp, prawn

Dendrobranchiata

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Charles D. Derby and Martin Thiel his work was clearly very impressive given the available technology and was a model for neuroethological research in that generation and beyond. Figure 1.2 presents a sampling of the work of Bethe. The techniques used for the first time by him on living organisms (microsurgical ablations) are still employed today in neuroethological studies. Combinations of these techniques

A

B

C

Fig. 1.2. Examples of the neuroethological experiments by Albrecht Bethe in the 1890s to identify brain regions and neurons responsible for behavior of the shore crab Carcinus maenas. (A) A drawing from Bethe of a partially dissected crab, showing the central nervous system, including the brain (Gehirn) and some of its neural connections to the anterior end of the animal, esophageal ganglia (Schlundganglion) on the circumesophageal connectives encircling the esophagus (Ösophagus), and the ventral nerve cord formed from the fused thoracic and abdominal ganglia. (B) Bethe’s drawing of a brain showing neuropils, nerve roots, and the morphology of individual neurons based on methylene blue staining. (C) Bethe made various ablations of the nervous system of live animals to examine the behavioral roles of specific brain regions and nerves. He made small holes in the crab’s exoskeleton and then used various tools to alter the animal’s nervous system. His tools included iridectomy scissors used to cut nerves, and knives fabricated from insect pins to remove cell bodies, cut tracts between neuropils, and other microdissections. After the dissections, Bethe sealed the holes in the exoskeleton with wax and then examined the animal’s behavior. Panel C shows the behavior of crabs after cutting of the right circumesophageal connective (rechts Schlundcommissur). Two days after nerve transection, these ablations caused crabs to walk in a circular pattern, either to the left- (Linksgang) or right-hand direction (Rechtsgang). Bethe confirmed the anatomical location of his ablations by examining the histology of the brains of animals after completion of behavioral experiments. Panels A and B are from Bethe (1897a), and Panel C is from Bethe (1897b).



Nervous Systems and Control of Behavior: An Introduction

with modern genomics tools such as RNA interference and optogenetics promise to be powerful approaches in the future. Neuroscientific research using crustaceans has continued since these early studies. In the early 20th century, the works of the Swedes Nils Holmgren (1916) and Bertil Hanström (1926, 1928) are notable. Periodically, there have been reviews on the subject. Ted Bullock and Adrian Horridge’s two-volume Structure and Function in the Nervous System of Invertebrates from 1965 provided a thorough synthesis that included but was not limited to crustaceans. In 1961 and 1962, an influential two-volume series limited to crustaceans appeared—The Physiology of Crustacea, edited by Talbot Waterman. Waterman’s volumes were the main review materials on the subject until 20 years later, with the publication of a 10-volume series The Biology of Crustacea, published in 1982 and 1983 with Dorothy Bliss as the editor-in-chief. Two of those volumes were on nervous systems: Volume 3, Neurobiology: Structure and Function, and Volume 4, Neural Integration and Behavior, edited by Harold Atwood and David Sandeman—both are contributors to the present volume! Since 1982, three other volumes on crustacean neuroscience were published, based on two conferences held on the subject. Two of these volumes are conference proceedings that contain numerous short reports on specialized research projects: Frontiers in Crustacean Neurobiology, and The Crustacean Nervous System, both edited by Konrad Wiese and published in 1990 and 2001 respectively. The third volume was a set of review papers that emerged on select topics from the second of those conferences: Crustacean Experimental Systems in Neurobiology, published in 2002 and also edited by Wiese. In 2012, Nicholas Strausfeld published Arthropod Brains: Evolution, Functional Elegance, and Historical Significance, an incredible book focusing on neuroanatomy that deals wonderfully with all of the subtopics in the title.

THIS VOLUME Nervous Systems and Control of Behavior, the third volume in the new series The Natural History of the Crustacea—focuses on the functional organization of crustacean nervous systems and how nervous systems produce behavior. This volume is intended to synthesize the state of the field in crustacean neurobiology. It is aimed at both crustacean researchers and neurobiologists studying other taxa. This volume has 20 chapters authored by internationally recognized experts in their disciplines. The book is divided into three sections that build progressively on each other. The first section, “Principles of the Nervous System,” provides descriptions of the basic organization of the nervous system of crustaceans, including sensory, integrative, and motor systems. It focuses on functional organization, including morphology and physiology. It presents a broadly phylogenetic perspective, as much as possible from what is known about the major groups of crustaceans, though necessarily focusing on the better-studied group, the malacostracans, and even more specifically, the decapods. The six chapters in the first section serve as a foundation for the more specialized topics in the rest of the volume. The second section, “The Senses and Sensory Ecology,” has five chapters, each focused on how a particular sensory system functions toward allowing crustaceans to solve problems facing them, such as acquisition of food, shelter, and mates, within the unique context of the animals’ specific natural environment. The third section, “Neural Control of Behavior,” has seven chapters that present examples of behavior of crustaceans, from the relatively simple (gastric movements) to the much more complex (walking, escape, navigation, social interactions, and memory and learning), and how the nervous system controls the production of these behaviors. A final chapter presents how crustaceans are used as models in studies of the nervous system: in genomics research and as teaching tools. The themes and chapters are represented in Fig. 1.3.

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Charles D. Derby and Martin Thiel Adult neurogenesis Synapses

m s ste Sy us vo

Chemoreception & 10 chemical ecology

er N of

le s

so en

11 Mechanoreception

ry ol Ec

12

y

og

1

Visual ecology

S nd sa

History

3

9

se en

2

5

Vision

eS

Sensory systems Brain form & function

8

6

Th

Modulation Motor & muscle systems 4

7

Pr in cip

6

Crustaceans as 20 teaching tools 19 Learning & memory Path integration & decision-making

Neural Control of Behavior 18

17 Social status

16

15

Magnetoreception

Digestion & the 13 stomatogastric ganglion 14 Posture & walking

Escape & tailflipping Biological rhythms

Fig. 1.3. A summary of the topics and chapters in this volume. The image of the crayfish is modified from Keim (1915).

Chapters treat their subjects in parallel format. First is an introduction that develops the general context of the topic and identifies the main issues treated in the contribution. The main text of each chapter then presents to the readers, in a concise fashion, the current status of knowledge and understanding of the subject. This is by necessity not a comprehensive treatment, in that only a limited number of crustacean groups, topics, and references can be included. The treatment is broadly comparative, meaning it not only treats crustaceans but also attempts to point out similarities and contrasts with other animal taxa. Each chapter then considers future directions, discussing the most important questions left unanswered and the most promising directions for future research—what might be the general content of the next volume. Each chapter ends with a summary and conclusions, the take-home messages that will be of greatest value to all the readers.

THEMES AND HIGHLIGHTS Research on the neuroscience of crustaceans can have different major aims. One aim is to understand mechanisms underlying evolutionary adaptations within a natural historical context. This approach takes advantage of the fact that crustaceans inhabit and are adapted to a diversity of environments. This approach is traditionally called comparative physiology, and more recently includes, depending on the focus, neuroethology (Zupanc 2004), sensory ecology (Dusenbery 1992), and neuroecology (Zimmer and Derby 2011). This volume has many chapters that highlight this aim. Adaptations in the sensory abilities of crustaceans are described in overview chapters on nervous and sensory systems (­chapter 2 by Sandeman and colleagues, c­ hapter 3 by Mellon) and in chapters dealing with each of the senses. These include sensory abilities that we humans have, including vision (­chapter 8 by Glantz, ­chapter 9 by Cronin and Feller), chemoreception (­chapter 10 by Derby and Weissburg), and mechanoreception (­chapter 11 by Lenz and Hartline, ­chapter 14 by Cattaert). Given the huge diversity of environments in which crustaceans live, it is not surprising that collectively their sensory systems respond to environmental stimuli far beyond the five traditional senses of humans and include detection of the



Nervous Systems and Control of Behavior: An Introduction

earth’s magnetic field (­chapter 12 by Lohmann and Ernst), and wavelengths of light that humans cannot detect (­chapter 8 by Glantz, ­chapter 9 by Cronin and Feller). This comparative neuroscientific approach has yielded detailed understanding of neural mechanisms underlying several behaviors of crustaceans; one behavior is tail flipping. This form of escape locomotion is one of the best-understood behaviors at the neural level and is a model for neuroethology (­chapter 15 by Krasne and colleagues). Also well-studied is the neural control of posture and walking (­chapter 14 by Cattaert), biological rhythms (­chapter 16 by Fanjul-Moles), and social status (­chapter 17 by Herberholz). New approaches using computer and neuromechanical simulations and hybrid systems, combining both an in vitro nervous system and a robotic or simulated body, are increasingly being used. These approaches can help us understand the interactions between central networks, sensory feedback, and mechanical properties of musculoskeletal components in organizing and regulation locomotor activity. Progress is being made in understanding even more complex behaviors, such as path integration and decision-making (­chapter 18 by Zeil and Hemmi) and learning and memory (­chapter 19 by Tomsic and Maldonado). The second major aim of neuroscientific research on crustaceans is to use them as model systems to understand basic processes and functions of nervous systems and their components. This approach has taught us much about how sensory cues are detected and processed by nervous systems, as outlined in several chapters in the second section. One of the best-studied systems in all of neuroscience for understanding principles of flexible circuits and the underlying mechanisms of neuromodulation—that is, using chemicals to modulate specific synapses such that neural circuits can have multiple states and outputs—is the stomatogastric system of crustaceans. How crustaceans do so much with this network of only a couple dozen of neurons is described in ­chapter 5 by Harris-Warrick and in ­chapter 13 by Marder and colleagues. Many crustaceans produce new neurons throughout their lives, and a comparative view of adult neurogenesis is presented in ­chapter 7 by Schmidt. Basic principles of crustacean motor systems and synapses are treated in ­chapter 4 by Atwood and ­chapter 6 by Lnenicka. In fact, the value of using crustaceans as models for exploring basic processes and principles of neural function makes them outstanding preparations for teaching basic concepts in neuroscience, as summarized in c­ hapter 20 by Johnson and colleagues. This volume highlights how much more we know about crustacean neuroscience than presented in past review volumes on the subject just 10 or 20 years ago. For example, we know that new neurons are born continuously in the brains of many adult crustaceans, as has also been discovered in some other animals. We know that axons of some crustaceans are myelinated for rapid neural transmission, previously thought to be a feature unique to vertebrates. Crustaceans can be sensitive to many environmental stimuli previously not appreciated, including geomagnetic cues and some wavelengths of light outside of the spectrum visible to humans. We begin to understand at a molecular level processes underlying learning, memory, and other forms of plasticity. We know that neural circuits can be highly flexible in their outputs and that this is due to modulation of basic processes of cells and synapses by neuroactive substances. We know some of the biochemical and molecular underpinnings of the formation and maintenance of social status. We appreciate much more than before the functional organization of many parts of the crustacean nervous system.

FUTURE OPPORTUNITIES AND CHALLENGES Crustaceans offer many opportunities for neuroscientific research. The complexity in their behavior and nervous systems provides much source material to examine. Coupled with advances in technology, this allows that many subjects and issues can be addressed as

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Charles D. Derby and Martin Thiel never before. The behavior of crustaceans can be complex, sophisticated, intricate, and, of course, fascinating. Watching animals, particularly in the field, can yield so many insights about their behavior, adaptations, and sensory world. It is often the inspiration and starting point for laboratory- and field-based experimental studies that dive deeper into mechanisms and consequences of behavior. Equipment for digitally capturing and analyzing videos has made it standard fare to study behaviors of all types including very fast behaviors, behaviors performed in the dark, and behavior in the field. From this basis in behavior, reductionist approaches can be used to examine in detail the mechanisms underlying these behaviors, or synthetic approaches can be used to examine higher-order ecological issues. For example, the fields of sensory ecology and neuroecology combine techniques spanning from biophysical and molecular properties of nerve and muscle cells to community-wide impacts of trophic interactions, to understand the consequences that neural mechanisms of sensory systems may have on ecological interactions (Dusenbery 1992, Zimmer and Derby 2011). These fields can help us understand principles controlling the organization of communities by examining the processes of information transfer, the ability of animals to respond to signals indicating the presence of competitors, mates, food, dwelling sites, and other resources, and the effect of subsequent responses on community members. Barnacles, for example, are a foundation species and dominant competitors for space in rocky intertidal habitats. These crustaceans use the same proteins—α 2 -macroglobulins—to build defensive shells and as chemical cues to attract settling conspecifics (Clare 2011), and these same molecules are also cues used by predators and competitors to locate barnacles (Ferrier et al. 2011). Such “molecules of keystone significance”—which are relatively rare, mediate a variety of interactions between organisms and inf luence the distribution and abundance of species, the transfer of energy across multiple trophic levels, and thus play significant roles in structuring ecosystems (Zimmer and Ferrer 2007)—may well play important organizing roles in other crustacean-driven communities, and sensory and neuroecological approaches will be required to characterize them. Opportunities to use modern molecular techniques on crustaceans are increasing, but a limitation at present is there are relatively few complete genomic sequences for crustacean species. This is in part due to the relatively large sizes of crustacean genomes (Gregory 2012, Jeffery 2012). Now that the first crustacean genome is sequenced—Daphnia pulex (Colbourne et  al. 2011)—more sequences are on the way. For example, the i5K Insect and Other Arthropod Genome Initiative (http://arthropodgenomes.org/wiki/i5K) planned to sequence genomes of 5,000 arthropods between 2010 and 2015. Of the first 130 species planned to be sequenced, seven are crustaceans: two decapods, one amphipod, and four isopods. This is a good start, but more is necessary to be able to more broadly use the molecular techniques such as RNA interference (Kato et al. 2011) and optogenetics (Deisseroth 2011) that will allow exploration of mechanisms in more detail. There has been incredible progress in the field of crustacean neurobiology since the work of Bethe and others in the 19th century. Yet despite this, we are still far from achieving Bethe’s goal of knowing a crustacean’s connectome and using it to understand the neuronal basis of all behaviors of that species. But there is also so much potential for natural history, neuroethological, and biomedical explorations into the nervous systems of crustaceans. It will be interesting to see, 20+ years from now and hence in a future volume on crustacean neuroscience, how far we have progressed in our understanding of neuroscience and crustaceans.

ACKNOWLEDGMENTS We thank Dr. Manfred Schmidt for sharing his knowledge of the history of crustacean neuroscience, translating and discussing German literature including the papers of Albrecht Bethe, and



Nervous Systems and Control of Behavior: An Introduction

providing helpful comments on our introductory chapter. We also thank Dr. Brian Mulloney for sharing the image in Fig 1.3 of the crayfish from Keim 1915.

REFERENCES Ahyong, S.T., J.K. Lowry, M. Alonso, R.N. Bamber, G.A. Boxshall, P. Castro, S. Gerken, G.S. Karaman, J.W. Goy, D.S. Jones, K. Meland, D.C. Rogers, and J. Svavarsson. 2011. Subphylum Crustacea Brünnich, 1772. Pages 165–191 in Z.-Q. Zhang, editor. Animal biodiversity: an outline of higher-level classification and survey of taxonomic richness. Zootaxa, Vol. 3148. Magnolia Press, Auckland, New Zealand. Atwood, H.L., and D.C. Sandeman, editors. 1982. The biology of Crustacea, Vol. 3, Neurobiology: structure and function. Academic Press, New York. Bethe, A. 1895. Studien über das Centralnervensystem von Carcinus maenas nebst Angaben über ein neues Verfahren von Methylenblaufixation. Archiv für mikroskopische Anatomie und Entwicklungsgeschichte 44:579–622. Bethe, A. 1897a. Das Centralnervensystem von Carcinus maenas. Ein anatomisch-physiologischer Versuch. I. Theil. I. Mittheilung. Archiv für mikroskopische Anatomie und Entwicklungsgeschichte 50:460–546. Bethe, A. 1897b. Das Centralnervensystem von Carcinus maenas. Ein anatomisch-physiologischer Versuch. I. Theil. I. Mittheilung. Archiv für mikroskopische Anatomie und Entwicklungsgeschichte 50:589–639. Bethe, A. 1898. Das Centralnervensystem von Carcinus maenas. Ein anatomisch-physiologischer Versuch. II. Theil. III. Mittheilung. Archiv für mikroskopische Anatomie und Entwicklungsgeschichte 51:382–452. Bracken, H.D., A. Toon, D.L. Felder, J.W. Martin, M. Finley, J. Rasmussen, F. Palero, and K.A. Crandall. 2009. The decapod tree of life: compiling the data and moving toward a consensus of decapod evolution. Arthropod Systematics and Phylogeny 67:99–116. Bullock, T.H., and G.A. Horridge. 1965. Structure and function in the nervous system of invertebrates. Vols. 1 and 2. Freeman, San Francisco London. Clare, A.S. 2011. Toward a characterization of the chemical cue to barnacle gregariousness. Pages 431–450 in T. Breithaupt and M. Thiel, editors. Chemical communication in crustaceans. Springer, New York. Colbourne, J.K., et al. 2011. The ecoresponsive genome of Daphnia pulex. Science 331:555–561. Deisseroth, K. 2011. Optogenetics. Nature Methods 8:26–29. Dusenbery, D.B. 1992. Sensory ecology: how organisms acquire and respond to information. Freeman, New York. Ferrier, G.F., S.J. Kim, J.A. Loo, C.A. Zimmer, and R.K. Zimmer. 2011. Sensory mechanisms driving community ecological interactions. Integrative and Comparative Biology 51, Supplement 1:E41. Freud, S. 1882. Über den Bau der Nervenfasern und Nervenzellen beim Flusskrebs. Sitzungsberichte der Kaiserlichen Akademie der Wissenschaften in Wien 85:9–46. Gregory, T.R. 2012. Animal genome size database. http://www.genomesize.com Hanström, B. 1926. Eine genetische Studie über die Augen und Sehzentren von Turbellarien, Anneliden und Arthropoden (Trilobiten, Xiphosuren, Eurypteriden, Arachnoiden, Myriapoden, Crustaceen und Insekten). Kungliga Svenska Vetenskapsakademiens Handlingar Ser. 3, Bd. 4, No. 1:1–176. Hanström, B. 1928. Vergleichende Anatomie des Nervensystems der wirbellosen Tiere unter Berücksichtigung seiner Funktion. Springer Verlag, Berlin. Holmgren, N. 1916. Zur vergleichenden Anatomie des Gehirns von Polychaeten, Onychophoren, Xiphosuran, Arachniden, Crustaceen, Myriapoden und Insekten. Vorstudien zu einer Phylogenie der Arthropoden. Kungliga Svenska Vetenskapsakademiens Handlingar Series 2, 56:1–303. Jeffery, N.W. 2012. The first genome size estimates for six species of krill (Malacostraca, Euphausiidea): large genomes at the north and south poles. Polar Biology 35:959–962.

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Charles D. Derby and Martin Thiel Kato, Y. Y. Shiga, K. Kobayashi, S. Takishita, H. Yamagata, T. Iguchi, and H. Watanabe. 2011. Development of an RNA interference method in the cladoceran crustacean Daphnia magma. Development Genes and Evolution 220:337–345. Keim, W. 1915. Das Nervensystem von Astacus fluviatilis. Ein Beitrag zur Morphologie der Dekapoden. Zeitschrift für wissenschaftliche Zoologie 113:485–545. Martin, J.W., and G.E. Davis. 2001. An updated classification of the recent Crustacea. Natural History Museum of Los Angeles County, Science Series 39. Martin, J.W., and G.E. Davis. 2007. Historical trends in crustacean systematics. Crustaceana 79:1347–1368. Porter, M.L., M. Perez-Losada, and K.A. Crandall. 2005. Model-based multi-locus estimation of decapod phylogeny and divergence times. Molecular Phylogenetics and Evolution 37:355–369. Regier, J.C., J.W. Shultz, A. Zwick, A. Hussey, B. Ball, R Wetzer, J.W. Martin, and C.W. Cunningham. 2010. Arthropod relationships revealed by phylogenomic analysis of nuclear protein-coding sequences. Nature 463:1079–1083. Retzius, G. 1890. Zur Kenntnis des Nervensystems der Crustaceen. Das Zentrale Nervensystem. Biologische Untersuchungen Neue Folge 1:1–50. Richter, S., and G. Scholtz. 2001. Phylogenetic analysis of the Malacostraca (Crustacea). Journal of Zoological Systematics and Evolutionary Research 39:113–136. Sandeman, D.C., and H.L. Atwood, editors. 1982. The biology of Crustacea, Vol. 4, Neural integration and behavior. Academic Press, New York. Scholtz, G., and S. Richter. 1995. Phylogenetic systematics of reptantian Decapoda (Malacostraca: Crustacea). Zoological Journal of the Linnean Society 113:289–328. Schram, F.S. 2012. Comments on crustacean biodiversity and disparity of body forms. Pages 1–33 in L. Watling and M. Thiel, editors. The natural history of Crustacea, Vol. 1, Functional morphology and diversity. Oxford University Press, New York. Strausfeld, N.J. 2012. Arthropod brains: evolution, functional elegance, and historical significance. Belknap Press of Harvard University Press, Cambridge. Tsang, L.M., K.Y. Ma, S.T. Ahyong, T.-Y. Chan, and K.H. Chu. 2008. Phylogeny of Decapoda using two nuclear protein-coding genes: origin and evolution of the Reptantia. Molecular Phylogenetics and Evolution 48:359–368. von Reumont, B.M., R.A. Jenner, M.A. Wills, E. Dell’ampio, G. Pass, I Ebersberger, B. Meyer, S. Koenemann, T.M. Iliffe, A. Stamatakis, O. Niehuis, K. Meusemann, and B. Misof. 2012. Pancrustacean phylogeny in the light of new phylogenomic data: support for Remipedia as the possible sister group of Hexapoda. Molecular Biology and Evolution 29:1031–1045. Waterman, T.H. 1960. The physiology of Crustacea, Vol. 1, Metabolism and growth. Academic Press, New York. Waterman, T.H. 1961. The physiology of Crustacea, Vol. 2, Sense organs, integration, and behavior. Academic Press, New York. Wiese, K. 2001. The crustacean nervous system. Springer-Verlag, Berlin Heidelberg New York. Wiese, K. 2002. Crustacean experimental systems in neurobiology. Springer-Verlag, Berlin Heidelberg New York. Wiese, K., W.-D. Krenz, J. Tautz, H. Reichert, and B. Mulloney. 1990. Frontiers in crustacean neurobiology. Birkhäuser Verlag, Basel. Wirkner, C.S., and S. Richter. 2010. Evolutionary morphology of the circulatory system in Peracarida (Malacostraca; Crustacea). Cladistics 26:143–167. Zimmer, R.K., and C.D. Derby. 2011. Neuroecology and the need for broader synthesis. Integrative and Comparative Biology 51:751–755. Zimmer, R.K., and R.P. Ferrer. 2007. Neuroecology, chemical defense, and the keystone species concept. Biological Bulletin 213:207–224. Zupanc, G.K.H. 2004. Behavioral neurobiology: an integrative approach. Oxford University Press, New York.

2 ADAPTIVE TRENDS IN MALACOSTRACAN BRAIN FORM AND FUNCTION RELATED TO BEHAVIOR

David C. Sandeman, Matthes Kenning, and Steffen Harzsch

Abstract In eumalacostracan brains, the sensory inputs from the various receptor systems distributed on the head appendages can be traced to their respective neuropils. The comparison of the brain “ground patterns” of a number of eumalacostracan species with that of a notional ancestral form is used here to explore the adaptive changes that have occurred during the evolution of the Eumalacostraca and that could be related to their adoption of particular habitats and lifestyles. We suggest that adaptations to habitat or lifestyle are mainly confined to receptor systems and within the organization of the primary sensory neuropils. Here the changes are often more quantitative than qualitative, although in some instances (e.g., the optic neuropils in stomatopods), neuropil structure can reflect the significant anatomical changes in the receptor organ. Eumalacostracan brains all contain neuropils that can be considered as “higher integrative centers” because they receive no direct inputs from primary sensory fibers and have no direct outputs to motor neurons. These centers are the hemiellipsoid bodies, the terminal medullae and the accessory lobes. At this level, both qualitative and quantitative differences between the species can be found, some of which have a phylogenetic basis (e.g., the appearance of the accessory lobes) while others are possibly related to behavioral adaptations associated with habitat or lifestyle (e.g., loss of the olfactory neuropils in desert isopods). The eumalacostracan brain therefore exhibits plasticity at both the quantitative and qualitative levels that matches the radiation of these animals across a wide range of habitats and the adoption of a variety of lifestyles.

INTRODUCTION This chapter is concerned with the eumalacostracan crustacean brain, its ontogeny, and the relative differences in neuropil organization and size that may represent adaptive changes to the different habitats and lifestyles of different species. 11

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David C. Sandeman, Matthes Kenning, and Steffen Harzsch Animals reveal their presence either by their movement or by chemical cues. They live in a hierarchy in which they prey on those smaller or less able to defend themselves, or are preyed on by those larger or more ferocious than themselves. To avoid being preyed on, animals need to keep still and mask their own chemical cues. To survive, they need to detect the chemical cues or motion of potential prey. Significant areas of the brains of most animals are therefore given over to processing the inputs from the visual, chemical, and mechanical receptor systems. In the brains of the malacostracans, these are situated in the optic lobes and protocerebrum (vision, eyes), deutocerebrum (chemical and mechanical, antenna 1 or first antenna), and tritocerebrum (chemical and mechanical, antenna 2 or second antenna). These functional divisions are blurred beyond the primary receptor projection areas by relays of second and higher order neurons. For example, responses to visual stimuli have been found in interneurons that project to deutocerebral neuropils (the accessory lobes in crayfish; Sandeman D. et al. 1995), and projections from the olfactory lobes are known to ascend to the lateral protocerebral neuropils (terminal medulla, Derby and Blaustein 1998; hemiellipsoid body, Mellon et al. 1992a, Wachowiak et al. 1996, Sullivan and Beltz 2001, 2005). Information concerning spatial orientation from statocysts is relayed to motor systems throughout the body (eyes, antennae, locomotory appendages, telson), and mechanoreceptive information from the tritocerebrum ascends to the deutocerebrum (Sandeman D. et al. 1995). The different modalities soon, therefore, become integrated into what is probably a diffuse neuronal “gestalt” spread throughout the brain. Nevertheless, it is still possible to distinguish between the proportional projections of the various sensory systems to different areas of the brain and draw some conclusions from this in terms of the ensuing behavioral patterns. Behavioral activity involves several levels of the central nervous system, and all except the simplest reflexes usually include the integrated input from many receptor systems. At the lowest level, actions require not much more than the basic framework of a coordinated motor system coupled to sensory feedbacks that regulate cyclic locomotory actions such as walking, running, hopping, swimming, and flying. Such feedback loops, coupled to specificallytuned sensory systems, can rapidly compensate for unexpected external disturbance and prevent animals from stumbling on the ground or falling out of the air. At this level, the behavior is to a certain extent “automatic” and the associated neural components can be described and understood in terms of a relatively limited set of sensory and motor pathways. While such systems may appear self-contained, compensatory actions in freely-moving animals are integrated with higher order systems that determine the onset or cessation of bodily movement and control its direction and velocity. The basic motor framework also encompasses more than just locomotion, and there exist motor patterns that are responsible for grooming, feeding, fighting, mating, and so on, all of which employ the same appendages used in the compensatory reflexes and which are voluntarily regulated by the animal. Hence, the separation of behavior patterns into specific levels of complexity is less helpful than viewing the range of activities as a continuum extending from reflexive escape reactions, through compensatory actions to complex behaviors, which may indeed be dominated by one particular sensory input but seldom rely entirely on that modality. The complexity of the various behavior patterns could be defined in terms of the number of separate sequential components involved, their duration, and a requirement for learned elements. The neural substrate that is involved at this “cognitive” level in the behavioral continuum can be expected to be represented in the central nervous systems of all animals and not just those that are recognized as belonging to “higher” forms. “Cognitive” has been defined as “the mental action or process of acquiring knowledge through thought, experience and the senses” (Oxford English Dictionary), an ability seemingly reserved for the higher mammals alone. However, in some long-term crustacean behavior patterns (recognition of individual antagonists, social interactions, extensive seasonal migration, homing), learning and memory, both short and long term, appear to play a significant role, and even if these do not qualify as truly “cognitive,” they come close to it.



Adaptive Trends in Malacostracan Brain Form and Function Related to Behavior

The literature on the nervous systems and behavior of the malacostracans is extensive and cannot be covered in a single chapter. We have therefore selected heavily, trusting that the examples we have chosen illustrate the structural plasticity found in the brains of malacostracans that accompanies their adaptive radiation within different habitats and their adoption of various lifestyles.

THE PHYLOGENY OF MALACOSTRACA The crustaceans are a large and diverse group of arthropods. They range in size from tiny copepods of less than a millimeter in length, to long-legged spider crabs with a leg span of up to 1.8 meters. They are found in habitats extending from benthic volcanic vents, through coastal, estuarine, and freshwater habitats, to terrestrial areas including deserts. An extensive fossil record suggests that most major lineages of the crustaceans probably arose and diversified during the Precambrian. These then underwent a long period of independent evolution, leading to significant diversity within and among the groups. Genetic and developmental lability led to pedomorphosis, character convergence, and reversal, making the selection of useful phylogenetic characters difficult (Spears and Abele 1997). While crustacean phylogeny is not the main issue in this chapter, a brief account of those groups that are included here and their relationship to the larger group that make up the Crustacea as a whole is relevant in spite of the difficulties mentioned above. We have relied on and slightly modified the phylogenies proposed by Scholtz and Richter (1995), Richter and Scholtz (2001), and Wirkner and Richter (2010). Members of the Malacostraca are familiar to many and include the Stomatopoda (mantis shrimps), Decapoda (spiny lobsters, lobsters, crayfish, hermit crabs, crabs), Anaspidacea (Tasmanian mountain shrimps), Euphausiacea (krill), and Peracarida (shrimps, amphipods, isopods). For this account, we selected members of the Decapoda (Stenopodidea, Achelata, Homarida, Astacida, Anomala, Brachyura), Peracarida (Isopoda), and Stomatopoda, all of which are included in the Eumalacostraca (Fig. 2.1).

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Fig. 2.1. The phylogeny of the Malacostraca (based on Scholtz and Richter 1995, Richter and Scholtz 2001, Wirkner and Richter 2010). Bold text denotes the taxa from which examples in this chapter are taken. Question mark denotes uncertainty about this affiliation.

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David C. Sandeman, Matthes Kenning, and Steffen Harzsch

THE MALACOSTRACAN BRAIN Ground Patterns An important feature of the Eumalacostraca is that their brains are “modular” and an anatomical segmentation is retained, allowing the sensory inputs from the various receptor systems distributed on the head appendages to be traced to their respective neuropils. The hypothetical “ground pattern” suggested for the decapods (Sandeman and Scholtz 1995) can be used as a template for other members of the Eumalacostraca (Stomatopoda, Syncarida, Euphausiacea, and Peracarida) because the same areas of neuropil can be identified in all of them. We use the terminology of Sandeman et al. (1992) with the exception of the optic ganglia. For these, we adopt the terminology of the homologous neuropils in insects, namely, lamina, medulla, lobula, and lobula plate. We use the ground patterns here to compare the differences between the various species, which may be related to their adoption of particular lifestyles or habitats. We employ a color code (Fig. 2.2, center plate) to indicate four regions of the malacostracan brains (optic ganglia, protocerebrum, deutocerebrum, tritocerebrum) to which the different neuropils can be assigned based on their segmental ontogeny (see “Embryonic Development of the Brain”) and identifiable subdivisions within these (Fig. 2.2, center plate). In the ground pattern of a notional malacostracan, the brain is longer than it is broad (Fig. 2.2A) and includes a discernible lobula plate, and all lateral, medial, and midline protocerebral neuropils. The deutocerebrum lacks an accessory lobe and has only a single cluster of small “globuli” cells associated with each olfactory lobe. The lateral antenna 1 neuropils are as large as the olfactory lobes and are subdivided. The tritocerebral antenna 2 neuropils are elongated and relatively large. The generalized ground pattern of the Decapoda differs little from the malacostracan pattern except for the appearance among the Pleocyemata (Caridea, Stenopodidea, Reptantia) of a second cluster of “globuli” cells associated with each olfactory lobe (Fig. 2.2B). Figure 2.2 provides a general reference to the ground patterns of all the species considered in relation to their adoption of particular habitats and lifestyles. Embryonic Development of the Brain Clues to homologous structures, important for the derivation of ground patterns and phylogenetic relationships, can often be obtained from the study of the embryological development of the organisms of interest. Immunocytological techniques have extended the classical morphological studies and provided definitive descriptions of segmentation in crayfish (Fig. 2.3A,B) (Scholtz 1995, 1997, Alwes and Scholtz 2006, Sintoni et al. 2007, Vilpoux et al. 2008) and the maturation of neurons in the brain that express certain neurotransmitters (reviews Beltz 1999, Spindler et al. 2000) (Fig. 2.3C). Developmental studies provide the following picture of brain segmentation in malacostracan crustaceans (Fig. 2.3D–F): 1. The ocular protocerebral region is associated with visual input from the median and compound eyes. The lateral protocerebrum is located in the eyestalks and comprises the terminal medulla and the hemiellipsoid body. 2. The median protocerebrum houses the central complex and the anterior and posterior median protocerebral neuropils. 3. The most proximal retinotopic neuropil, the lobula, derives ontogenetically from the terminal medulla and so is part of the lateral protocerebrum. The development of the lobula

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Fig. 2.2. The ground and brain patterns of malacostracans. The subdivisions of the proto-, deuto-, and tritocerebrum can be identified by their shape and gray shade (see color version of this figure in center plate) provided in the key. (A) Malacostraca. (B) Stomatopoda. (C) Decapoda. (D) Stenopus hispidus. (E) Birgus latro. (F) Eureptantia. (G) Isopoda. Abbreviations: Optic ganglia: L, lamina; M, medulla; LO, lobula; LOp, lobula plate. Protocerebrum: HBN (HBN1, HBN2), hemiellipsoid body (subdivisions thereof); TM, terminal medulla; AMPN, anterior median protocerebral neuropil; PMPN, posterior median protocerebral neuropil; PBr, protocerebral bridge; CB, central body; LL, lateral lobe. Deutocerebrum: OL, olfactory lobe; LAN, lateral antenna 1 neuropil; OGTN, olfactory globular tract neuropil; AL, accessory lobe; MAN, median antenna 1 neuropil. Tritocerebrum: AnN, antenna 2 neuropil; TN, tegumentary neuropil; Receptors, chiasmata, and tracts: R, retina; CH1, first optic chiasm; CH2, second optic chiasm; OGT, olfactory globular tract.

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Fig. 2.3. (A) Embryo of the marbled crayfish around 45% of embryonic development labeled with a general nuclear marker (from Seitz et al. 2005, with permission from John Wiley & Sons). (B) Engrailed immunoreactivity in an embryo of the marbled crayfish around 40% of embryonic development (from Sintoni et al. 2007, with permission from Springer) in which the labeled cells indicate the boundaries of the individual segments. (C) Developing brain in an embryo of the marbled crayfish slightly before 40% of development labeled with a probe against actin (from Vilpoux et al. 2006, with permission from Springer). (D) Schematic, idealized model of the egg-nauplius brain of crayfish, which, it is proposed here, conserves information on the possible appearance of the brain architecture of an early crustacean or even arthropod ancestor. The dotted lines depict the outlines of the egg-nauplius. A question mark denotes uncertainty about the connection. The four neuromeres of the naupliar brain are arranged in mirror symmetry across the horizontal center of the stomodeum (from Vilpoux et al. 2006, with permission from Springer). (E) Whole mount of lobster embryos Homarus americanus labeled against synaptic proteins (synapsin) to reveal the embryonic nervous system (Harzsch, Benton, Beltz unpublished, with permission from authors). (F) Development of the optic lobes in an embryo (E75%) of the crayfish Cherax destructor labeled with phalloidin, a probe against actin (from Vilpoux et al. 2006, with permission from Springer). Abbreviations: A1, 2, antenna 1 and 2 anlagen; aMD, anterior portion of the mandibular neuromere; AnN, antenna 2 neuropil; Ca, Cp, anterior and posterior pre-esophageal commissures; CP, caudal papilla; DC, anlage of deutocerebrum; HBN, hemiellipsoid body anlage; ION/SON, inferior and superior esophageal nerve; L, lamina; La, anlage of labrum; LO, lobula; LPC, lateral protocerebrum; M1, M2, anlagen of maxilla 1 and 2; MD, anlage of mandible and mandibular neuromere; MDC, mandibular commissure; ME, medulla; MPC, medial protocerebrum; OA, optic anlagen; OL, olfactory lobe; PC, protocerebrum; POC, post-esophageal commissure; PT, protocerebral tract; R, developing retina; ST, stomodeum; T1-4, anlagen of thoracic limbs 1 to 4; TC, anlage of tritocerebrum; TM, terminal medulla. Scale bars: A, 100 µm; B, 25 µm; C, 75 µm; E, 100 µm; F, 20 µm.



Adaptive Trends in Malacostracan Brain Form and Function Related to Behavior

plate is intimately linked to the lobula (Strausfeld 2005) and hence it too can be regarded to be part of the lateral protocerebrum. On the other hand, the first two retinotopic neuropils, the lamina and medulla, are protocerebral but seem to develop independently of the lateral protocerebrum. 4. Neuropils associated with the antenna 1 input such as the olfactory and accessory lobes and the medial and lateral antenna 1 neuropils are deutocerebral. 5. Neuropils associated with antenna 2 such as the antenna 2 neuropil are tritocerebral as is the tegumentary neuropil. The commissural ganglion has tritocerebral components as well as contributions from the mandibular neuromere. Traditionally, the esophagus has been thought to pierce the developing brain between the deuto- and tritocerebrum (Fig. 2.3C). However, recent developmental data revealing the presence of deutocerebral fibers behind the esophagus suggest that, as in pterygote insects, the esophagus actually pierces the deutocerebrum (Fig. 2.3D) (Vilpoux et al. 2006).

ADAPTIVE CHANGES RELATED TO BEHAVIOR Complex behavioral patterns involve many receptor systems, but often one particular modality may dominate. Predatory animals, for example, frequently depend on their visual systems to detect and then pursue their quarries. This does not mean that the chemosensory or mechanoreceptive systems have no part to play, and quite often the different receptor systems will be employed in sequence during the behavioral procedure such as the use of tactile information during the actual grasping of the prey. Habitat and lifestyle can play a significant role in determining the dominant receptor system. Predatory crustaceans, such as mantis shrimp, that live in conditions of high illumination may be expected to have capitalized on the visual system. Others, such as the fiddler crabs living in tidal mudflats whose dominant aspects of behavior (mating, feeding) take place in air, may have adopted visual systems that are particularly suited to the visual environment in which they live. Alternatively, if they are nocturnal or live in conditions of little or no light such as cave crayfish, they may have either very poorly developed eyes or none at all. Terrestrial crustaceans, such as land crabs, may have modified olfactory systems to operate in air or, like desert-living isopods, not have them at all and rely instead on contact chemoreception. Alternatively, animals that operate without good visual systems, or like spiny lobsters and Australian freshwater crayfish that are mainly active in conditions of poor light, or live in turbid water, can compensate with highly sensitive mechano-chemoreceptive appendages. Examples that illustrate the above and are included here are grouped together under the subsections of visual, chemosensory, and exploratory mechano-chemosensory behavior. Visual Behavior Overview Motion is perceived by an eye when an image moves across the photoreceptive cells in the retina. This can result from the movement of an object in front of the stationary eye or by the movement of the eye past a stationary object. As long as animals remain stationary, any motion in the visual field can be confidently assumed to be extrinsic to itself. During voluntary movements however, the situation is more complex because it is then necessary to distinguish between self-induced and extrinsic image movements. A strategy to confront this problem—development of eyes that can move independently of the body—has evolved several times. This allows an image to be

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David C. Sandeman, Matthes Kenning, and Steffen Harzsch stabilized, within limits, onto one area of the eye. Motion of small targets within this stabilized visual field is more easily detected than within an entire visual field that is also moving over the retina. A further development is found in active predators in which specific areas of the eyes, often with enhanced optical resolution, are fixated on an object of interest. In both cases, the strategy requires mounting the eyes in the head so that they can move in both vertical and horizontal planes, and the development of a neuromuscular system that can precisely control the eye movements in relation to the image on the retina. Many species of crustaceans have stalked compound eyes, a highly-developed motor system that controls the eye movements, and a well-developed set of compensatory reflexes that are driven by the eyes themselves, the balance organs, and the proprioceptors in the legs. Amongst the malacostracans, stalked eyes are a characteristic feature of the stomatopods, decapods, all euphausids, and syncarids (except Bathynellacea, all of which are blind). The eyes of the peracarids however are sessile. As examples of adaptations of visual systems related to behavior, we selected the stomatopods, fiddler crabs, and cave crayfish, and limited ourselves to comparisons of the anatomy of the optic neuropils (see also ­chapters 3, 8, 9, 18, and 19 in this volume). Habitat and Lifestyle Stomatopods The four extant superfamilies of the Stomatopoda are the Squillidae, Lysiosquillidae, Gonodactylidae, and Bathysquillidae. All are marine, and members of the first three families live in water up to a depth of 300 m (Caldwell and Dingle 1975). The lysiosquilloid and gonodactyloid species predominate in brightly illuminated shallower water, and the squilloid species in the deeper areas. The bathysquilloids include the only species to extend their habitat beyond the 1,000 m mark (Marshall et al. 2007). Stomatopods are characterized by the development of the second maxillipeds as raptorial appendages that can be very rapidly extended to either crush (smashers) or impale (spearers) their prey. The gonodactyloids have adopted the smashing strategy and also tend to inhabit cavities in reefs from which they emerge to prey on reef organisms. Among the stomatopods, they occupy the greatest diversity of habitats. The squilloids and lysiosquilloids construct burrows in the soft substrata in which they live and have less armor than the gonodactyloid species (Caldwell and Dingle 1975). Lysiosquilloids lie at the entrance of their burrows with only the eyes and antenna 1 protruding through holes in a small mucus trapdoor over the entrance to the burrow. Small fish or other prey are “speared” as they pass by (Mead and Caldwell 2011). The stomatopods exhibit complex behavior patterns in establishing their burrows or cavities, mating, and food capture. Those that live in shallower waters depend heavily on their visual systems, both for the recognition of conspecifics and prey items (Caldwell and Dingle 1975), but their olfactory and tactile senses are also important in some contexts. The eye movements of the stomatopods differ from the bilaterally-coupled motions of other stalk-eyed malacostracans; instead, each eye is able to move, chameleon-like, independently of the other and exhibits a high degree of freedom about all three rotational axes. They are almost continually in motion (Horridge 1978, Land et al. 1990) and will track rapidly moving targets (Cronin et al. 1988). The eye muscle systems that control these complex movements (Jones 1994) are unlike those known in the crabs (Burrows and Horridge 1968) or crayfish (Mellon 1977). The eyes of the stomatopods are unusual among the malacostracans in that each eye is subdivided into three anatomically distinct regions in which a narrow midband of ommatidia separates two outer hemispheres. In line with the phylogeny of the stomatopods, the midband contains six rows of ommatidia in both the gonodactyloids and lysiosquilloids, two in the



Adaptive Trends in Malacostracan Brain Form and Function Related to Behavior

squilloids and none in the bathysquilloids. The midband contains 14 of the 16 different types of photoreceptors that are found in the eye (see Marshall et al. 2007 for review). The photoreceptor types in the hemispheres are like those seen in other malacostracans (Stowe 1980). Behavioral evidence indicates that at least the gonodactyloids and lysiosquilloids have both color and polarization sensitivity (Marshall et al. 1996, 1999). The tripartite retina of the stomatopods provides the optic ganglia with separate streams of information; the hemispheres, spatial and monocular stereoscopic information; the midband, color, ultraviolet light, and the plane of polarized light (Marshall et al. 2007). Fiddler Crabs Fiddler crabs inhabit tidal expanses of sand or mud and so live in a flat world. They dig burrows in the substratum, take refuge in these during high tides, and emerge at low tide to feed on the detritus that accumulates and remains in the surface layers of the mud or sand after the tide has receded. Some species live as mixed-age and -gender communities in relatively confined areas where an individual carries out its normal activities in a space that may not be more than a square meter. For the study of behavioral interactions of animals within a community, the fiddler crabs offer significant advantages and these have been exploited by many researchers (see Zeil and Hemmi 2006, and c­ hapter 18 in this volume). The lives of fiddler crabs are centered on their burrows, from which they make excursions. This exposes them to threats from predatory birds and from conspecifics that can gain access and possession of their burrows. Recognition of these threats and the response to them (i.e., immediate return to the burrow) depend entirely on the visual system, and an examination of their eyes reveals features that suit them perfectly for the role they play. The eyestalks of the fiddler crabs are long and are folded laterally into grooves in the carapace when the animals enter their burrows. On emergence from the burrows, the eyestalks are extended vertically upward and are positioned close together over the midline of the body. The ommatidial surfaces of the eyes are located at the distal end of eyestalks and distributed around the surface of the cylindrical eyestalk. When extended, the two eyes together include a visual field of 360º in the horizontal plane. The corneal surface in the axial plane of the eyestalk, on the other hand, is almost straight. The vertical visual field is therefore a relatively narrow window that corresponds to an area just above and below the horizon. Many ommatidia are directed toward the horizon, which has the consequence of increasing the resolving power of the eye in this region. This is a useful feature when it is considered that it is in precisely this region of high contrast between the ground and the sky that objects larger than the animals themselves could appear and perhaps represent a predatory threat. Anything below the crab’s horizon is either the same size or smaller than itself and so not a predatory threat, but possibly a conspecific rival (Zeil et al. 1986, 1989, see also ­chapter 18 in this volume).

Blind Crayfish Several species of crayfish have invaded caves in recent geologic history and occupy many limestone cave systems in regions in North America. Among these are the obligate cave-dwelling (troglobitic) Procambarus erythrops (Mellon 1977) and Orconectes australis packardi, and the facultative cave species Cambarus tenebrosus (Cooper et al. 2001). In caves or in aquaria in the laboratory, both O. australis packardi and C. tenebrosus sweep the long second antennae across the area in front of them while walking, implying a significant dependence on mechanical and chemical information. Recordings of the changes in heart rate indicate the sensitivity of cave crayfish to white light (but not to infrared or dim red light), waterborne vibrations, and chemical cues, although they may show no overt reaction to such stimuli

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David C. Sandeman, Matthes Kenning, and Steffen Harzsch (Li et al. 2000). The aversive reaction of O. australis packardi to a broadband light source (Li and Cooper 1999) may be mediated by photoreceptors situated in the caudal ganglion (Larimer 1966) or in the median protocerebrum of the brain (Page 1982, Sandeman et al. 1990). Perhaps one of the most unexpected responses of both P. erythrops and O. australis packardi is their compensatory eye movements evoked by subjection to angular rotation about their long axes (roll) or when their bodies are held stationary while the substratum on which they are standing is rotated about the vertical axis (yaw). Recordings from motor nerves and eye muscles in P. erythrops and Procambarus clarkii (Mellon and Lnenicka 1980) show that these muscle systems are not only active but produce the correct compensatory response of the eyestalks. In sighted P. clarkii, these compensatory actions stabilize the image of the environment on the retina. The reflexes would appear redundant in the blind P. erythrops but are retained. The function is not clear. The protective reflex withdrawal of the eyestalk into its socket is also retained in P. erythrops (Mellon 1977). Visual stimuli that evoke the behavioral “startle response” (raising the chelae into a defensive posture) in P. clarkii are ineffective in the other species, implying that they are unable to detect the change in light intensity that occurs when a hand is passed over the top of the animal, or that the stimulus has no meaning for the animals because small electrical responses can be recorded even from the cornea of P. erythrops following abrupt changes in light intensity (Mellon 1977) (see also ­chapters 8 and 9 in this volume). When compared with the surface-living P. clarkii, the cornea and underlying retina of cave crayfish exhibit different degrees of reduction in the size of the eyestalk and retina and the presence of dark pigment behind the cornea. The eyestalk of C. tenebrosus is about half the size of P. clarkii but has a retinal surface that covers proportionately about the same surface area as that of P. clarkii (Cooper et al. 2001). The eyestalks of P. erythrops are cone-shaped with a much reduced retinal surface located at the apex of the cone. The cornea of C. tenebrosus is faceted, indicating the position of underlying ommatidia, whereas there are no facets on the cornea of P. erythrops (Mellon 1977). The eyes of O. australis packardi are the most reduced of the three species mentioned here, being mere stubs with no corneal surface or pigmentation (Cooper et al. 2001). Central Neural Pathways and the Ground Pattern The retinotopic projection pattern of photoreceptor axons of the ommatidia in the three different regions of the tripartite stomatopod eye is retained in the optic neuropils. Midband cartridge diameters are larger than those from the hemispheres and their projections are anatomically distinguishable as a separate area (accessory lobe) in the medulla and lobula (Kleinlogel et al. 2003). Although these central neural changes are related to the adaptation of the visual system for the complex predatory and social behaviors of the stomatopods, they are superimposed on the basic malacostracan ground pattern and appear to be confined solely to the optic ganglia (Fig. 2.2, compare A with B). The photoreceptor system on the other hand contains features that are unique to the stomatopods (Marshall et al. 2007). While not as extensive as in the stomatopods, the adaptive changes in fiddler crabs are also predominantly confined to the receptor system and limited essentially to the elongation of the eyestalks, the particular distribution and size of the ommatidia over the retina, and the shape of the eye surfaces. The location, size, and gross appearance of the optic neuropils are little different from that of the basic ground pattern. Despite a complex social lifestyle, a uniquely adapted visual system, and a specialized habitat, the central nervous system ground pattern appears to have been conserved just as it has in the predatory mantis shrimp, which could hardly be more different in terms of habitat and lifestyle from the grazing fiddler crabs.



Adaptive Trends in Malacostracan Brain Form and Function Related to Behavior

Blind crayfish provide an opportunity to deduce which neuropils are essential for vision and, because they have retained those in the lateral protocerebrum, also those important for chemoor mechanosensory inputs. The optic neuropils are reduced in size in both P. erythrops and O. australis packardi, and have lost the laminar organization that characterizes these neuropils in sighted crayfish (Mellon 1977, Cooper et al. 2001). While both species exhibit some sensitivity to light, they do not have a visual sense that includes the perception of motion or the formation of an image. However, despite this loss of a functional visual system, not only does P. erythrops retain an intact eye muscle system virtually identical to that of P. clarkii, but also the motor neurons that drive these muscles respond to nonvisual inputs that evoke oculomotor reflexes (Mellon 1977). The loss of the visual input has not extended to the motor system although the eye movements that are produced, and which in a seeing animal would serve to stabilize the visual image on the retina, are clearly redundant. Strong connections via the large, fine-fibered olfactory globular tract that extends from the olfactory and accessory lobes (when present) to the hemiellipsoid body and to the terminal medulla are a common feature of the malacostracan ground plan. Despite the significant reduction of the optic neuropils in O. australis packardi, the numbers of axons in the projection of the olfactory globular tract to the eyestalk in these animals is greater than that of both the sighted P. clarkii and C. tenebrosus (Cooper et al. 2001). Here, it would seem that olfaction has become more relevant for the blind animals, and the hemiellipsoid body and terminal medulla are retained. The remainder of the brains of these animals has not been carefully investigated but most likely show little deviation from the eureptantian ground pattern. Chemosensory Behavior Overview Chemoreception is perhaps the most important and most widely exploited sensory modality in the malacostracans, and even those animals with exceptional visual systems, the stomatopods, have highly-developed chemosensory receptor systems. A wide range of behaviors depend on the reception of chemical stimuli, and our current knowledge on this subject has been comprehensively reviewed in the multi-author volume Chemical Communication in Crustaceans edited by Breithaupt and Thiel (2011). Chemical stimuli are used in every aspect of the lives of crustaceans, and in many cases the behavior patterns are complex and of long duration involving a sequence of actions that depend on other senses (mechanoreceptive and visual) (Hay 2011). Chemoreception in the malacostracans may be separated into two classes: “Olfaction” refers to the detection of odorants that are present in the surrounding medium. “Distributed chemoreception” refers to chemosensitive sensilla that are bimodal and respond to both chemical and mechanical stimuli (Schmidt and Mellon 2011, see ­chapters 3 and 10 in this volume). In terms of possible adaptive changes in the chemoreceptive systems of the malacostracans, we confine ourselves here to a special set of chemoreceptive sensilla, the aesthetascs and their olfactory receptor neurons (ORNs), that are located on the first antennae, and the area within the brain to which these receptors project, the olfactory lobes. Olfactory information, like that of other sensory inputs, is relayed beyond the primary sensory neuropils to higher order centers in the brains of the malacostracans, and these projections are considered elsewhere in this chapter. As examples, we selected freshwater crayfish, coconut crabs, and woodlice (see also ­chapters 3 and 10 in this volume).

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David C. Sandeman, Matthes Kenning, and Steffen Harzsch Habitat and Lifestyle Crayfish Freshwater crayfish are found worldwide in ponds and streams and have been favorite experimental animals for neurobiologists for many decades. While the crayfish olfactory system clearly plays a part in finding nourishment, it plays a central role in mating behavior, including the associated establishment and recognition of dominance hierarchies. The chemicals involved in mating and dominance contests are contained in the urine released from sphincter-controlled pores (Bushmann and Atema 1996) on the basal segment of the second antennae, and directed in water currents toward the conspecific receiver. These currents are generated by movements of the scaphognathite (gill bailer), maxillipeds, and fan organs associated with the mouthparts (Breithaupt 2001). The generation of water currents to disperse chemicals is critical in those species that live in stagnant water where diffusion is very slow. Both reproductively active male and receptive females of Orconectes rusticus and Procambarus leniusculus release pheromones during pairing (Berry and Breithaupt 2010). Agonistic behavior in freshwater crayfish and the establishment of dominance has been comprehensively reviewed by Moore (2007) and by Herberholz in ­chapter 17 of this volume. Such contests between males begin with ritualized, non-damaging behaviors but can escalate to levels in which participants lose appendages. A characteristic of the behavior is the excretion by both contestants of pheromone-bearing urine. If the same animals are matched repeatedly, the fight durations will decrease and a stable dominance hierarchy will be established. The olfactory recognition of social status is a critical factor in the establishment and maintenance of the dominance hierarchy (see Breithaupt 2011 for review). In the malacostracans, the detection of soluble or volatile odorants in solution or in air and at a distance from the source is carried out by the specialized olfactory sensilla on the first antennae, the aesthetascs (Hallberg et al. 1992, Hallberg and Skog 2011), and also by bimodal chemo-mechanoreceptive sensilla (e.g., hooded sensilla: Cate and Derby 2002), which are also responsive to chemicals that arrive in solution. The sensilla are probable homologs of those found broadly distributed on both the first antennae and body surfaces of the Palinuridae, Scyllaridae, and Nephropidae (Cate and Derby 2002, see also ­chapters 3 and 10 in this volume). The structure of the aesthetascs in several species of freshwater crayfish is known (Tierney et al. 1986, Sandeman and Sandeman 1996) and conforms to the general malacostracan pattern (Hallberg et al. 1992, Hallberg and Skog 2011) in that they have a relatively thin-walled cuticle and up to several hundred chemosensory cells surrounded by enveloping cells. Each sensory cell has dendrites from which transformed cilia arise. In addition, the cuticle of the aesthetascs in spiny lobsters may act as a molecular sieve, selecting for molecules having a molecular weight of less than 8.5 kDa (Derby et al. 1997). Coconut Crabs Although the large coconut crabs, Birgus latro, of the remote Indian and Pacific ocean islands have long amazed biologists by their size and terrestrial adaptation, the details of their behavior and in particular their emphasis on aerial olfaction has only recently received focused attention (see Drew et al. 2010 for review), and much is still unknown. Nevertheless as a terrestrially adapted species, they are evidently opportunistic hunters and scavengers that are normally solitary but can be found in aggregations at sites that contain an abundant supply of a favored food source. Copulation, during which a spermatophore is transferred to the female and deposited close to her gonopores (she has no seminal receptacle), takes place out of water as does the extrusion and fertilization of the eggs by the female (Drew et al. 2010). The egg mass is held beneath the



Adaptive Trends in Malacostracan Brain Form and Function Related to Behavior

abdomen of the female until shortly before hatching. At this time, the egg-bearing females move from their shoreline shelters out onto areas (intertidal flats or rocky ledges) that are washed by waves. Here the females cling to the substratum and release their eggs as the waves wash over them. Aerial olfaction is clearly important for these animals. They are able to detect and find sources of odor and are particularly sensitive to oligosulfide compounds that are emitted from decaying meat (Stensmyr et al. 2005, Krieger et al. 2010, Hansson et al. 2011). The use of pheromones in intraspecific communication does not appear to have been studied, although it may well be used by males to find females during the mating period. Initial tracking studies point to the existence of a relatively restricted home range (600 m) in some animals, although individuals can undertake longer sorties (1600 m) (Hansson et al. unpublished data cited in Drew et al. 2010). Even with the little information that we have on B. latro, here is a long-lived organism (estimated life span up to 40 years, Fletcher as cited in Drew et al. 2010) that exhibits a refined and olfactorily-oriented feeding behavior and a migratory-based mating procedure that depends on a guided and goal-oriented performance. The volume of the brain areas that are related to these behaviors is large in comparison with species with simpler lifestyles and more restricted habitats (Krieger et al. 2010). The thin-walled aesthetascs of marine malacostracans are unsuited to terrestrial conditions because they would most likely dry out after prolonged exposure to air. Nevertheless, the coconut crab B. latro has retained both the aesthetasc sensilla and the basic olfactory system of the aquatic malacostracans although with some significant modifications. At the receptor level, the aesthetasc sensilla are present along the terminal portion of the lateral flagellum of the first antenna. They are short and scale-like and packed in rows along the ventral surface (which is directed forward in an active animal). The individual aesthetasc sensilla are also asymmetrical: the cuticle covering the exposed surface is thin, that of the unexposed surface, much thicker (Stensmyr et al. 2005, Hansson et al. 2011). Differences in diffusion coefficients of volatiles in air and chemicals in solution can also play a part in the design of aesthetasc sensilla in aquatic or land-living Crustacea (Mellon and Reidenbach 2011). Modifications within the aesthetasc sensilla, such as the enclosure of the basal bodies and ciliary segments within a lymph space deep within the flagellum, are features that represent a convergence with the insects (Stensmyr et al. 2005). That the aesthetascs of the coconut crabs are sensitive to volatile odors and particularly to those that are also attractive to insects that feed on carrion has been convincingly demonstrated with behavioral and physiological techniques (Stensmyr et al. 2005, Hansson et al. 2011). Wood Lice The isopods occupy a wide range of habitats from marine (Idotea baltica), littoral (Ligia oceanica), terrestrial (Porcellio scaber, Armadillium vulgare), and desert (Hemilepistus reaumuri) (Harzsch et al. 2011). In terms of the degree of terrestrialness, isopods have achieved the highest level, defined as “fully terrestrial and able to conduct all biological activities on land.” Here they surpass even the land crabs, whose early larval stages require a marine environment. The desert-living H. reaumuri is monogamous, lives in burrows, and has a remarkable kin recognition behavior based on specific chemical information, or badges. Family members are recognized according to the nature of a unique, non-volatile, chemical badge produced by the individual and combined with that of the family members and contained in the cuticle. When these animals molt, the badge is temporarily lost but it is regained by remaining in close contact with the family members within the burrows (Linsenmaier 1987, 2007; see Thiel 2011 for review). The isopods provide a good example of the changes that can take place during the transition from marine to terrestrial habitats; the marine species such as I. baltica possess aesthetasc

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David C. Sandeman, Matthes Kenning, and Steffen Harzsch sensilla, albeit fewer in number than in the decapods (Harzsch et  al. 2011). However, unlike the coconut crabs, the terrestrial isopods P. scaber, A. vulgare, and H. reaumuri have dispensed with the malacostracan olfactory system. Their first antennae are much reduced and the second antennae appear to have assumed dominance as sensory structures (Harzsch et al. 2011). In particular, and of relevance for the kin-recognition in H. reaumuri, is the presence of an apical sensory cone at the tip of the second antenna that is used to touch the cuticle of conspecifics and so identify the chemical signature signaling family or stranger (Linsenmaier 1987, 2007). The lack of aesthetasc sensilla does not, however, prevent some terrestrial isopods from detecting volatile, airborne odors (Thiel 2011). Central Neural Pathways and the Ground Pattern The aesthetasc sensilla on the first antennae in all the malacostracans are characterized by their exclusive and unilateral projections to the olfactory lobes, which are paired, usually spherical neuropils in the lateral deutocerebrum. The lobe contains cone-shaped columns of synaptic regions that radiate out from the center of the lobe and that are the target of the primary sensory axons of the aesthetasc sensilla (Sandeman and Denburg 1976). The regions between the columns, or glomeruli, are generally devoid of synapses (Sandeman and Luff 1973, Blaustein et al. 1988, Sandeman et al. 1992, Mellon and Alones 1993, Schmidt and Ache 1997; reviews: Schachtner et al. 2005, Schmidt and Mellon 2011, and ­chapter 10 in this volume). The total volume of the olfactory lobe, the volume occupied by the glomeruli, their number, and the number of aesthetasc sensilla that project to the glomeruli were all found to vary in a study of 17 decapod species (Beltz et al. 2003). The data show a continuum across the groups from those that favor a high convergence of ORNs from many aesthetascs onto few glomeruli, to those where ORNs from fewer aesthetascs are shared among many glomeruli. While not diagnostic, it would seem that those species that have the lowest number of sensilla per glomerulus are often found in estuarine, intertidal habitats. Intermediate ratios are found in several freshwater crayfish and marine crabs, whereas the highest ratios are found in the large marine clawed and spiny lobsters. Hence in terms of habitat, and perhaps also lifestyle, there appears to be an interaction between the receptor input and the central neuropil to which the olfactory receptor neurons project (Fig. 2.4). The olfactory lobes of the crayfish most closely resemble those found in the eureptantian ground pattern (Sandeman et al. 1992, Sandeman and Scholtz 1995) (Fig. 2.2F). The olfactory lobe of the coconut crab is significantly modified, not only in that it has three instead of two subdivisions of the lobe but also because the associated accessory lobe is greatly reduced and the protocerebral hemiellipsoid body expanded (Fig. 2.2E) (Krieger et al. 2010). The olfactory lobe in the coconut crab is also very large, occupying about 40% of the total brain neuropil volume and has over 1,000 glomeruli. In terms of the number of interneurons in the olfactory pathway, the coconut crab surpasses the insects (Krieger et al. 2010). This may not, however, be a feature related to the adaptation of the system to the detection and analysis of airborne odors because the neuropil volume, glomerular number and volume, the aesthetasc count, and the ratio of the number of aesthetascs per glomeruli in B. latro are very similar to that of the marine Panulirus interruptus (Table 2.1). The olfactory glomeruli of B. latro and Coenobita clypeatus are sharply divided into cap, subcap, and base regions, and these coincide with a regionalization of the hemiellipsoid bodies into a cap, core 1, and core 2 (Harzsch and Hansson 2008, Krieger et al. 2010). The isopods, like the cave crayfish, provide us with an example of the consequence at the central level of the loss of a specific receptor organ. Olfactory lobes can be identified in the brains of I. baltica by their spherical shape and glomerular neuropils (Fig. 2.2G), whereas these

7

Habitat: Estuarine Freshwater Terrestrial Marine

0

9

Jasus edwardsii

8

Panulirus interruptus

7

Birgus latro

6

Percnon planissimum

5

Panulirus argus

4

Cancer borealis

3

Libinia dubia

Procambarus clarkii

1

Coenobita clypeatus

Uca minax

2

2

Cherax destructor

Uca pugilator

1

3

Petrolisthes coccnicus

Callianassa australiensis

4

Cherax quadricarinatus

5

Sesarma sp.

(Aesthetasc count/glomeruli #)

6

Homarus americanus

Adaptive Trends in Malacostracan Brain Form and Function Related to Behavior

Uca pugnax



10 11 12 13 14 15 16 17 18

Fig. 2.4. The ratio between the number of aesthetasc sensilla on the flagellum of antenna 1 and the number of glomeruli in the olfactory lobe reveals a tendency for the species to cluster into groups related to habitat: Species 1 to 5, estuarine; 6 to 8 and 11, freshwater; 9 and 15, terrestrial; 10, 12 to 14, and 16 to 18, marine (based on data from Beltz et al. 2003 and Krieger et al. 2010).

are absent from the brains of the terrestrial P. scaber, A. vulgare, and H. reaumuri (Harzsch et al. 2011). These isopods do not appear to be limited in their behavior in any way, having adapted sensilla on the second antenna to serve as both contact and olfactory sensilla (Thiel 2011). Also, while the large olfactory lobes (but perhaps more likely, the expansion of the hemiellipsoid bodies) may provide the coconut crabs with a neural substrate for olfactory memory, H. reaumuri is clearly able to learn and distinguish the chemical badges of its kin with no olfactory lobe at all. In summary, as seen in the visual system, adaptations to habitat and lifestyle would appear to be predominantly reflected in the receptor organs, and alterations in these are accompanied by central modifications involving the expansion or loss of elements of the original ground pattern. Exploratory Mechano-Chemosensory Behavior Overview Many malacostracan species are active in the dim light of evening or early morning or at night, or live in water that is often turbid. Crustaceans employ their first and second antennae, mouthparts, and legs in the directed exploration of their surroundings. Many of the sensilla on these appendages are sensitive to both mechanical and chemical stimuli and hence, despite the active nature of this behavior, it is not purely “tactile” in the sense of the active touch or haptic behavior found in vertebrates. Nevertheless, it is likely that crustaceans are able to discern the spatial and chemical nature of their immediate environment by such means. The anterior mechanosensory organs of the spiny lobsters and some crayfish are primarily their second antennae, particularly in cave crayfish but also in some epigean species, such as the Australian freshwater crayfish Cherax destructor, that are nocturnally active and frequently found in muddy or turbid ponds and streams. In spiny lobsters, the second antennae also play a defensive role, being relatively stiff with the basal segments covered in strong sharp spines. In some malacostracan species, particularly those with a diurnal lifestyle and that depend on

25

Brachyura

Anomura

Thalassinida

Homarida Astacida

Achelata

Taxon

153,833 280,103 18,947 229,666 39,338 58,705 14,887 17,779 13,14 8,034

19,585 28,041

9,790,377 6,588,788 120,352,292 374,681,700 12,359,013 165,730,818 20,327,317 28,765,244 6,617,077 4,558,497 3,114,604 3,012,080

287,884 117,862 616,475 591,583 110,975 74,298

344,922,004 154,068,687 591,956,438 141,159,589 24,187,019 24,735,814

Panulirus interruptus Panulirus argus Jasus edwardsii Homarus americanus Cherax destructor Cherax quadricarinatus Procambarus clarkii Callianassa australiensis Coenobita clypeatus Birgus latro Petrolisthes coccineus Cancer borealis Libinia dubia Percnon planissimum Sesarma sp. Uca minax Uca pugilator Uca pugnax

Glomerular volume µm³

ON neuropil volume µm³

Species

799 1,338 655 733 454 495 446 284 234 374

503 235

1,202 1,332 961 249 230 334

Glomeruli number

519 1700 328 540 319 555 33 39 28 26

133 22

1,786 1,255 1,537 1,262 130 237

Aesthetasc count

0.650 1.270 0.501 0.737 0.703 1.121 0.074 0.137 0.120 0.070

0.264 0.094

Convergence ratio (aesthetasc count/ glomeruli #) 1.486 0.942 1.599 5.068 0.565 0.710

Table 2.1  Interspecific morphometric differences. Comparative data relating neuropil and glomerular ­volumes, glomerular numbers, and the ratios of aesthetasc sensilla to glomerular number across a range of reptantian species (redrawn from Beltz et al. 2003, with permission from Wiley)



Adaptive Trends in Malacostracan Brain Form and Function Related to Behavior

vision, such as the stomatopods and rock crabs that inhabit rocky shores, the second antennae are small. In the stomatopods, however, it is the first antennae that play an important mechano-chemosensory role in mating behavior and that are coupled with the olfactory sense in the detection of pheromones (Mead and Caldwell 2011). The cleaner shrimp Stenopus hispidus has very long first and second antennae. The examples we consider here are the Australian freshwater crayfish, gonodactyloid stomatopods, and a stenopid cleaner shrimp (see also c­ hapter 3 in this volume). Habitat and Lifestyle Crayfish Cherax destructor, an Australian freshwater crayfish, is nocturnally active and forages on plants, other arthropods, and carrion. It is solitary and lives in complex underwater burrows that it defends and can use to survive periods of drought (Olszewski 1980, Reynolds 1980). Individuals have a home range that they patrol nightly, at times leaving the water to forage on the banks (Reynolds 1980). Given the relatively complex underwater environment, and the establishment of a home burrow, it is more than likely that the animals have a good spatial knowledge of their near environment and that they use mechano-chemosensory information to find their way around in the dark and at times muddy waters. Cherax destructor, if placed overnight with fish in an aquarium, will capture and eat these (Sandeman unpublished observation). Given that capture took place in low light conditions, one might assume mechano-chemosensory mediated behavior although some crustaceans and other arthropods have highly sensitive eyes that can operate in extremely low light intensities (see ­chapters 8 and 9 in this volume), so that vision cannot be totally ruled out in the above example. On the other hand, Euastacus spinifer, another Australian freshwater crayfish, actively hunts for tadpoles by antennating the substratum and not relying on vision (Turvey and Merrick 1997). The visual system in C. destructor appears to play a less important part in the search for stationary food items; both sighted and blinded crayfish stimulated by food odor will immediately begin to search the area, antennating the area around and in front of them. Provided with a small novel object, such as a polystyrene ball, both sighted and blinded animals will attack and attempt to eat this but only if they come across it with antennae, chelae, or legs (Zeil et al. 1985). Touching the second antennae of sightless C. destructor results in the animal lunging rapidly toward the point of contact and striking with its chelae. The direction and distance covered in such attacks are correlated with the angle of the antenna at the moment of contact and the point of contact along the antennal flagellum (Zeil et al. 1985, Sandeman and Varju 1988). Crayfish will also direct their second antennae at the source of a distant mechanical disturbance (Tautz et al. 1981, Masters et al. 1982, Tautz 1987). Like many other animals, when C. destructor is placed in a novel environment, it will explore. In an aquarium containing objects, blindfolded animals walk along the walls and stop to explore any object they come across with the second antennae. Placed in an arena empty of objects, blindfolded animals walk around the walls, “trailing” the nearside antenna along the wall. The antenna is flexible and “rolls” partially back past the animal as it trails the flagellum along a surface (Sandeman 1989). After a series of four repeated exposures to an empty arena, the period of exploration decreases, signifying habituation. An immediate increase in the exploratory activity (i.e., dishabituation) follows the introduction of short partitions projecting from the side walls of the arena. That such dishabituation also follows the removal or repositioning of the partitions implies that the animals are able to detect changes in the topography of their environment using mechano-chemosensory inputs alone and retain this information for at least 24

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David C. Sandeman, Matthes Kenning, and Steffen Harzsch h. Animals with immobilized second antennae no longer respond to topographical changes in the arena (Basil and Sandeman 2000). Part of the mechano-chemosensory exploratory strategy of C. destructor apparently depends on a bilateral comparison between the information received by the two antennae because in a T maze, animals with one immobilized or denervated antenna turn more often toward the untreated antenna (McMahon et al. 2005). The basal segment of the second antenna in crayfish is formed from the fusion of basipodite, ischiopodite, and meropodite. The flagellum of the second antenna is borne on the next proximal segment, the carpopodite, and is devoid of muscular tissue. The articulation of the basal segments is monitored by chordotonal organs. The same anatomical organization occurs in spiny lobsters (Rossi-Durand and Vedel 1982) and American lobsters (Sigvardt 1977). In C. destructor, the articulation of the basal segments of the second antenna is such that the flagellum can be directed at any point in space within a solid angle in the horizontal plane from about 10º on the contralateral side to 170º on the ipsilateral side. The excursion in the vertical plane extends from 10º below the horizontal in front of the animal to almost 180º at the back (Sandeman 1985). The second antennae in C. destructor are almost as long as its body, a feature shared with spiny lobsters and troglobitic crayfish. Mechanoreceptors on the second antennae of several species of crayfish have been described (Astacus leptodactylus: Tautz et al. 1981, Bender et al. 1984; Procambarus spp.: Taylor 1968, 1975a, 1975b) and are known to be highly sensitive to water vibration (Tautz et al. 1981, Masters et al. 1982). Four different types of setae were identified in scanning electron micrographs on the flagellum of the second antenna in C. destructor. These are the following: 1. Feathered procumbent setae that project across the arthrodial membrane linking the segments of the flagellum. These do not appear to be innervated in either C. destructor (Sandeman 1989) or in A. leptodactylus (Bender et al. 1984). 2. Short smooth setae that project at right angles to the flagellum and that comprise several subtypes. They resist deflection toward the base of the flagellum but hinge easily forward to lie flush against the flagellum. 3. Long smooth setae with their bases contained in large sockets. 4. Short stubby peglike setae (Sandeman 1989). Similar setal types with the exception of the peg setae occur on the second antennae of both Homarus americanus (Tazaki 1977) and A. leptodactylus (Tautz et al. 1981). The long smooth setae in C. destructor are highly mobile and can be observed to move with the slightest disturbance of the bathing medium. Electrophysiological measurements showed that responses to the motion of the bathing medium are abolished if all the long smooth setae are plucked off (Sandeman 1989). The short smooth setae are relatively insensitive to water motion, but respond with rapid phasic bursts when deflected. Stomatopods In stark contrast to the crayfish, the second antennae of the stomatopods play a very different role. The flagellum (endopodite) of the second antenna is relatively short. The exopodite is modified into a large elongated oval-shaped scale that in some animals is highly colored and used in agonistic displays. Such displays involve raising the cephalothorax, extending the first antennae and the scales of the second antennae, and the spreading and extension of the raptorial meri (Caldwell and Dingle 1975). Mating in these animals involves a complex process in which the males approach the females at the entrance to their burrows. Chemical signals are sent by both male and female individuals and driven toward each other with circular propulsive movements of the maxillipeds.



Adaptive Trends in Malacostracan Brain Form and Function Related to Behavior

The male also investigates the female using the first antennae (Mead and Caldwell 2011), possibly to detect both distant and contact pheromones with the aesthetascs and bimodal mechano-chemoreceptors. The behavioral observations of the stomatopods would suggest that the first antennae have assumed the mechano-chemosensory function of the second antennae of the decapods and play an important role in both mating behavior (Mead and Caldwell 2011) and prey capture (Schaller 1953). Cleaner Shrimps Cleaner shrimps are found in the Caridea (Urocaridella spp., Periclimenes spp., Lysmata spp.) and in the Stenopodidea (Stenopus spp.). Although belonging to three separate subfamilies within these taxa, they share some significant common anatomical and behavioral features. All are colored and have both red and either white or yellow bands or spots on their bodies and legs, although in many species the rest of the body is transparent. All have long, flexible first and second antennae, and in Lysmata amboiensis and Stenopus hispidus the first antennae are particularly noticeable, the two rami being almost as long as the second antennae. Organisms that live by cleaning others have to advertise their services in some way and avoid being taken as prey. Cleaner fish have a characteristically banded body coloration and undertake “dances” in which they swim up and down in the water column over their cleaning stations which are visited by much larger client fish (Grutter 2004). Cleaner shrimps also use this combination of repeated oscillatory motion coupled with color banding. Individuals of Urocaridella spp. signal their presence and readiness to clean by rocking their bodies back and forth. Stenopus spp. and Periclimenes spp. employ this strategy and in addition whip their antennae back and forth when potential clients are nearby (Becker et al. 2005). The relatively long first and second antennae and the emphasis placed on the coloring and movement of these in the unique behavior of these animals would suggest some discernible adaptive changes within the brain. There do not appear to be any available data on the nature of the sensilla on either the first or second antennae of these animals. Central Neural Pathways and Ground Patterns In the malacostracan ground pattern, axons from the second antennae project to the antenna 2 neuropils in the tritocerebrum, and this is the pattern in the crayfish, stomatopods, and cleaner shrimp S. hispidus (Fig. 2.2D). Despite the apparent dependence of the crayfish C. destructor on the input from the second antennae, the antenna 2 neuropils are not particularly large. They do, however, exhibit a striated appearance in histological sections caused by fiber bundles that run across the longitudinal axis of the neuropil, although this is not as pronounced as that found in Penaeus monodon and Pagurus bernhardus with their very long second antennae (Sandeman et al. 1993, Krieger et al. 2012) or in S. hispidus (Sandeman unpublished observation). While not confirmed, it is conceivable that the striations within the neuropil represent an ordered spatial projection from mechanoreceptors along the length of the flagellum. Unlike the stomatopods and the cleaner shrimps, the crayfish have large accessory lobes that receive projections from the antenna 2 neuropils (Sandeman D. et al. 1995). In the stomatopods, the first antennae have become dominant mechano-chemosensory organs, while not losing their olfactory role provided by the aesthetascs. The projection of the axons from the first antennae conforms to the malacostracan ground pattern: aesthetasc sensilla project exclusively to the olfactory lobes while all the other types end in the lateral antenna 1 neuropil in the medial deutocerebrum. As in other malacostracans, this is not a complete separation between chemoreception and mechanoreception because bimodal receptors that are

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David C. Sandeman, Matthes Kenning, and Steffen Harzsch both chemoreceptive and mechanoreceptive project to the lateral antenna 1 neuropil. The aesthetasc/olfactory lobe system mainly mediates complex behaviors associated with the detection of intraspecific chemicals such as recognition of dominance and kin and in mating behaviors (Schmidt and Mellon 2011, ­chapter 10 in this volume). The lateral antenna 1 neuropil in the stomatopods is large and bilobed (Fig. 2.2C). The bilobed nature of the lateral antenna 1 neuropil is also a feature of the malacostracan ground pattern; however it is more pronounced in the stomatopods. In addition, the stomatopod lateral antenna 1 neuropils exhibit a pronounced pattern of fiber bundles running transversely to the long axis of the neuropils. The elongated first and second antennae of the stenopid cleaner shrimp S.  hispidus is reflected in both the anatomy and volume of neuropil that is dedicated to these inputs (Fig. 2.2D). The bilobed structure of the lateral antenna 1 neuropils is even more pronounced than in the stomatopods, the antenna 2 neuropil is elongated as in the penaeids, and both the lateral antenna 1 neuropil and antenna 2 neuropil have the pattern of fiber bundles running transversely to the long axes of the neuropils. S. hispidus has a well-developed olfactory lobe; however the combined volume of the lateral antenna 1 neuropil and the antenna 2 neuropil, measured from silver impregnated serial sections, is nine times larger than the olfactory lobe (Sandeman, unpublished observation). Given the nature of cleaner shrimp behavior, the large increase in the volume of the lateral antenna 1 neuropil in comparison with the malacostracan ground pattern (Fig. 2.2, compare A with F) may reflect an increase in the number and importance of mechanoreceptive and bimodal mechano-chemoreceptors. The adaptive tendencies in the brains of the species we have examined that are related to exploratory mechanosensory behavior are, like those related to vision and olfaction, largely reflected in the volume, and perhaps complexity, of the neuropils that are associated with modifications to the sensory appendages, be these eyes, olfactory systems or mechano-chemosensory appendages. The malacostracan ground pattern can be recognized in all cases, with the significant exception of the appearance of the accessory lobes in the brains of the eureptantians (Fig. 2.2, compare A with G).

MULTIMODAL INTEGRATIVE BEHAVIOR The cognitive ability of mammals involves the cortex, an area of the brain with very large numbers of neurons, which receives no direct connections from primary sensory neurons, nor any direct outputs to muscles. It is a “higher” center by virtue of its separation from the “basic” framework that we describe in the Introduction, by relays of interneurons that convey partially processed and coded information that is “multimodal” in that it comes from many sensory systems. Are such higher centers represented in the brains of invertebrates, and if so, do these exhibit adaptations that are related to their behavior? Long-term and complex behavior patterns appear to be a characteristic of the larger reptantian crustaceans. Among their ranks, we find species that are both the largest and longest living of all arthropods. Although size in itself is not a guarantee for a large brain containing many neurons (e.g., the dinosaurs), at least there is space for a large brain should it be developed. While some arthropods with small brains (honeybees, ants, fruit flies) do indeed carry out complex behavior patterns, large brain size in terms of neuron numbers appears to be associated with a wider variety and flexibility of complex behavior patterns, cephalopod molluscs providing a good example. Longevity is not necessarily a feature that requires cognitive abilities, an example being the cephalopod mollusk Octopus vulgaris, which has a life span of only 12–15 months (Katsanevakis



Adaptive Trends in Malacostracan Brain Form and Function Related to Behavior

and Verriopoulos 2006). Nevertheless an extended life span could well be an important adaptive feature for lobsters that live much longer, providing them with the potential to reproduce over many years and to learn from experience. Their situation is very different from that facing ephemeral mayflies with an adult life span between a few minutes to several days, depending on the species. The adults of these insects have no functional mouthparts and once eclosed, their role is solely to mate, and then they die (Peters and Campbell 1991). Complex Behavior Patterns There is no shortage of examples among the malacostracans of complex behavior patterns related to finding nourishment, shelter, territory defense, mating, kin recognition, and brood care. Nevertheless, an area that seems to require a little more than these basic life requirements is that of social interactions among gregarious species during homing, the occupation of common shelters, cooperative behavior during seasonal migration, and the establishment of dominance hierarchies (see also ­c hapter 17 in this volume). It is difficult to conceive such behavior patterns being performed without elements of learning and memory. The spiny lobsters are perhaps the best studied members of the malacostracans in this regard and provide us with some good examples of long-term, complex behavior patterns. Two examples from the spiny lobster Panulirus argus are reviewed here; these are the orientation and homing behavior and the complex social behavior involving communal defensive tactics and offshore seasonal migrations. Orientation and Homing The spiny lobsters P. argus spend the day in shelters, or dens, in reefs. At night, they leave their dens to forage in open areas hundreds of meters away from the reefs, returning before daybreak to their reef and often to the same dens (Herrnkind and McLean 1971). Early tagging studies showed that two lobsters released at distances of 457 m and 3218 m away from their capture site were recaptured at the original site after 4 and 6 days, respectively (Creaser and Travis 1950). Displacement and sonic tracking studies of lobsters carried to about 200 m from their capture site on the reef confirmed the ability of even blinded lobsters to orient accurately toward their home sites (Herrnkind and McLean 1971). Reef-dwelling spiny lobsters forage widely over the areas surrounding their reef dens and will choose those areas containing the richest source of nourishment, which includes among others, molluscs, arthropods, echinoderms, sponges, and polychaetes (Cox et  al. 1997). Reef dens during nonmigratory periods can contain one to several lobsters and are seldom full. Animals within the dens will direct their spiny second antennae toward the entrance. Spiny lobsters do not have offensive appendages such as the large chelipeds of the clawed lobsters and crabs, or the raptorial maxillipeds of the stomatopods. Instead, when approached by a predatory trigger fish or a diver’s hand, they will antennate (whip) the intruder with the flagella of the second antennae and then lunge forward to drive off the threat with the strong spines around its base (Atema and Cobb 1980). When a number of lobsters reside together in the same den, they will all direct their antennae toward an intruder and thus present a formidable and thorny barrier (Herrnkind et al. 2001, Briones-Fourzán et al. 2006). Spiny lobsters also stridulate when threatened or attacked by predators (Atema and Cobb 1980). The path taken by spiny lobsters to their foraging areas and back to their reef dens is not random (Herrnkind et al. 1975, Jernakoff 1987). Spiny lobsters can detect and will orient in relation to the direction of water movement (surge) caused by wave action (Herrnkind and McLean 1971). They are also sensitive to geomagnetic information and walking animals will deviate

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David C. Sandeman, Matthes Kenning, and Steffen Harzsch from their chosen course when submitted to artificially generated fields that reverse the natural horizontal, but not vertical, geomagnetic field. Spiny lobsters therefore possess a magnetic field detection system sensitive to field polarity (Lohmann et al. 1995, Boles and Lohmann 2003, ­chapter  12 in this volume). Nevertheless, in addition to these broad orientational strategies, spiny lobsters returning from foraging areas may also use learned local structural features such as the edges of erosional undercuts along sea grass beds (Cox et al. 1997). Complex Social Behavior Adult spiny lobsters are gregarious and tend to aggregate. They are attracted over some distance to conspecifics by chemical cues (Zimmer-Faust et al. 1985) and to dens from which such cues emanate (Zimmer-Faust and Spanier 1987, Childress and Herrnkind 2001). Hence the final stages of homing, at least for those denning with conspecifics, is aided by chemical cues provided by others already in residence (Childress and Herrnkind 2001, Horner et al. 2008). Alternatively, lobsters that are physically damaged through attacks from predators exude chemicals that appear to act as warning cues since approaching lobsters will avoid dens containing such conspecifics (Shabani et al. 2008, Briones-Fourzán et al. 2008). Panulirus argus is also renowned for the extensive offshore migrations that it undertakes in autumn and that are characterized by a unique “queuing” behavior that has been likened to the flocks that birds form during their long migratory flights. Even in captivity, spiny lobsters have a propensity to line up behind other moving individuals, and the behavior can be seen in animals that return from open nocturnal feeding grounds to their diurnal shelters in the reefs. At the end of the summer, animals in captivity are seen to increase their activity and in circular tanks will form queues and walk along one behind the other for hours. Again this increase in activity has been likened to the “Zugunruhe” that is exhibited by migratory birds shortly before they initiate their migrations. The behavior of spiny lobsters during their migratory period differs markedly from that seen in nonmigratory periods. Codenning becomes much more common, with some dens completely filled to overf lowing. Migratory movements off the Bahamas and the Florida east coast occur during the day (Herrnkind and McLean 1971). Unlike the generally solitary foraging sorties made by nonmigratory spiny lobsters, migrating animals very seldom leave their dens alone. Instead they form “queues” in which the animals line up one behind the other in single file with the second antennae directed at an angle of about 45 degrees lateral to the midline. Constant contact with the individual ahead is maintained with either the inner f lagella of the first antennae, anterior walking legs, or second antennae (Herrnkind and McLean 1971). Migratory queues form in front of the dens when three to five individuals line up and set off together along the migratory pathway. These are followed by other queues that become longer as more individuals join until the den empties, resulting in a number of separate queues all heading in the same direction. Migrating lobsters move away from their protective reefs and over exposed areas of sand and are subject to predation from diurnally active triggerfish. To counter this threat, they exhibit some remarkable defensive behaviors that rely on cooperation between the individuals in the queue. Attacking trigger fish are first confronted with whipping and directed second antennae, followed by the lobsters in the queue forming a “rosette,” the entire queue adopting a circular formation from which the second antennae all point outward. Shorter queues that are attacked break from the single file and form a phalanx in which the lobsters form up side by side, facing the threat (Herrnkind et al. 2001). Migrating lobsters will rest both in areas of reef that lie in their path and in open areas. In reefs, they will aggregate in available crevices and if these fill up, cluster around the entrance.



Adaptive Trends in Malacostracan Brain Form and Function Related to Behavior

In open areas, rosette formation appears to be the usual defensive strategy that is adopted (see Herrnkind et al. 2001 for review). The social, homing, and migratory behaviors of P. argus and other species of spiny lobsters are some of the most complex found in the malacostracans. Depending on visual, olfactory, tactile, and geomagnetic senses, these animals provide a good example of multimodal integrative behavior, and the expectation that their central nervous systems may exhibit some areas beyond the optic ganglia, olfactory lobes, and antenna 2 neuropils to which the primary sensory neurons project. Higher Integrative Centers Three neuropil areas in the crustacean brains that are candidates for higher integrative centers can be recognized across species and contain interneurons responding to the selective stimulation of several different sensory systems. These three neuropil areas are the terminal medullae, the hemiellipsoid bodies, and the accessory lobes. To avoid confusion here, the hemiellipsoid body is treated as a separate neuropil and not included as a subsection of the terminal medulla as in some earlier studies (Blaustein et al. 1988). These areas qualify as “higher order” in that they receive information exclusively from second or higher order neurons, contain a large number of small fibered neurons, and have no primary sensory or motor neuron arborizations within them. The hemiellipsoid bodies and terminal medullae are part of the lateral protocerebrum and often, but not always, located in the eyestalks of those species that have them. There is a strong connection between the olfactory and accessory lobes and the lateral protocerebrum via the large olfactory globular tract. These connections are summarized in Figure 2.5. The terminal medullae, hemiellipsoid bodies, and accessory lobes are notable for their complexity and the amount of brain space that has been devoted to them, suggesting roles that go beyond simple reflexive behavior and that may involve more sophisticated processing related to orientation within the environment during homing or migration, recognition of suitable mating partners, and social interactions. All of these require aspects of learning and memory (see ­chapter 19 in this volume). Terminal Medulla The terminal medulla is a collection of smaller neuropil areas that are partly confluent with one another but that can be anatomically distinguished along their peripheral borders into a series of lobes. The neuropil structure within these areas is tangled, has a heterogeneous appearance, and contains both coarse and fine fibers. As such it resembles many other areas in the brain. There are eleven neuropil areas in the terminal medulla in both Procambarus clarkii and Panulirus argus, excluding the two that make up the hemiellipsoid body (Blaustein et al. 1988). Tracts between some terminal medulla neuropils and the optic ganglia, and from others to the olfactory and accessory lobes, suggest that the terminal medulla is a center in which a considerable amount of interaction between the olfactory and visual input occurs. This is supported by physiological studies in which electrical recording from, and labeling of, individual neurons that project to neuropils within the terminal medulla. Those with somata in the trito-, deuto-, or median protocerebrum and projections to the terminal medulla exhibit sensitivities to stimuli that range from unimodal chemical, through bimodal chemical and mechanical, to multimodal including chemical, mechanical, and photic. Neurons with their somata in the terminal medulla and that had morphologically complex branching were excited by chemicals applied to the first antennae but inhibited by tactile or visual stimuli (Derby and Blaustein 1988). The terminal medulla of B. latro is, like in all the other species so far examined, not geometrically arranged into columns or layers and is connected to the optic neuropils through the axons

33

A

B

C

E

F

D

Fig. 2.5. (A and B) Brain of the giant robber crab Birgus latro (Anomura; from Krieger et al. 2010, with permission from BioMed Central). (A) A single vibratome section (100 μm) double labeled against allatostatin-like immunoreactivity (AST, left) and synapsin immunoreactivity (SYN, right) to highlight the hemiellipsoid body (HBN), terminal medulla (TM), olfactory lobe (OL), accessory lobe (AL), and lateral antenna 1 neuropil (LAN) (B) A silver-impregnated microtome section (10 μm) showing the same neuropil areas as in A and the location of somata clusters 5, 6, 9, and 11. (C and D) Marine hermit crab Pagurus bernhardus (Anomura). (E and F). Shore crab Carcinus maenas (Brachyura). Immunolocalization of RF-amide-like neuropeptides on vibratome sections of the brains (from Harzsch et  al. 2011, with permission from Elsevier). (C)  P.  bernhardus, right olfactory lobe (OL; anterior is toward the top) with olfactory glomeruli (OG) and antenna 2 neuropil (AnN). Insets: higher magnification of olfactory glomeruli showing RF-amide-like (left) and synapsin-like immunoreactivity (right) to demonstrate their subdivision into cap (C), subcap (SC) and base (B). (D) P. bernhardus, transverse segmentation (arrows) of the antenna 2 neuropil. (E) C. maenas, horizontal section showing an overview over the brain. (F). C. maenas, higher magnification of the right olfactory lobe (OL) to show the radial arrangement of the cone-shaped glomeruli (OG). C–F from Harzsch et al. 2011, with permission from Elsevier. Scale bars: A, 500 µm; B, 100 µm; C, 200 µm; insets, 25 µm; D, 200 µm; E, 500 µm, F, 100 µm.



Adaptive Trends in Malacostracan Brain Form and Function Related to Behavior

in the optic tract. The medial area of the terminal medulla receives projections from the lobula and the lobula plate which are ontogenetically derived from the terminal medulla and so may be considered to be part of the lateral protocerebrum rather than being optic neuropils. The terminal medulla of B. latro also receives a significant deutocerebral input from the olfactory lobe via the axons in the olfactory globular tract (Krieger et al. 2010). Hemiellipsoid Bodies The hemiellipsoid bodies are separated into two areas in the spiny lobsters (I and II, Blaustein et al. 1988), the crayfish P. clarkii and O. rusticus (neuropil I and II, Sullivan and Beltz 2001), the crayfish C.  destructor (HBI and HBII, Sullivan and Beltz 2005), and the clawed lobster Homarus americanus (core and cap neuropil, Sullivan and Beltz 2001). While anatomically probably homologous, there are some differences in structure between the crayfish and the lobsters and also differences in the areas that are targeted by the axons of the projection neurons that ascend from the deutocerebrum. In crayfish, the hemiellipsoid bodies are targeted by projection neurons from the accessory lobes and not from the olfactory lobes (Mellon et al. 1992a, 1992b, Sullivan and Beltz 2001, 2005), whereas those from the olfactory lobes target primarily the terminal medullae. There is therefore very little overlap of the input from these two areas at the level of the lateral protocerebrum. Furthermore, in the crayfish C. destructor, the projection neurons from the cortex and medulla of the accessory lobes project separately to the inner (HBII) or outer (HBI) neuropils of the hemiellipsoid bodies. In the clawed lobster H. americanus, however, while there is still a pronounced separation of the projections from the accessory lobes and the olfactory lobes to the cap neuropil (I) and the core neuropil (II), the cap neuropil also receives branches from the olfactory lobes that sweep over the outer surface of the cap and form a layer within the cap (Sullivan and Beltz 2001). This is interesting in terms of the situation that is found in the hemiellipsoid bodies of the coconut crab B. latro, as described later in this section. Electrophysiological and anatomical investigation of the hemiellipsoid bodies in the crayfish C. destructor and P. clarkii brought a class of interneurons to light that are local to the hemiellipsoid bodies and that branch extensively in the two neuropil areas of the hemiellipsoid bodies (Mellon et al. 1992a, 1992b, Mellon and Alones 1997, Mellon 2000, McKinzie et al. 2003). There are about 200 of these so-called parasol cells that exhibit various patterns of continuous activity that are modified by olfactory, tactile, and visual stimuli (Mellon and Alones 1997, Mellon and Wheeler 1999, Mellon 2000, 2003) (Fig. 2.6). The somata of these cells are located ventral to the terminal medulla, and their primary neurites project to and arborize within either the HBI or HBII neuropils (McKinzie et al. 2003). The parasol cells, despite the common location of their somata, can therefore be considered to represent two classes of neurons depending on the nature of their inputs. In addition to the dendritic projection of the parasol cells in either of the HB neuropils, an axon extends to arborize in the terminal medulla (McKinzie et al. 2003). The hemiellipsoid bodies in B. latro are large, located medially in the brain and not in the eyestalks, and separated into three distinct areas, reflecting to some extent the situation in H.  americanus (Sullivan and Beltz 2001). The structure of the hemiellipsoid body in B.  latro resembles the layers of an onion (Krieger et al. 2010), with a peripheral and hemispherical cap neuropil enclosing two neuropil areas CO1 and CO2, which may be homologous with the neuropil areas I and II of the crayfish, or, depending on the status of the cap neuropil in the lobster may represent a subdivision in B. latro or the H. americanus core neuropil. At present, details of the pathways between these areas in B. latro and the deutocerebral neuropils that could indicate homologies are not yet available. Such information would indeed be of interest in relation to the changing functions of the olfactory lobe and hemiellipsoid body that may have compensated for the absence of large accessory lobes in B. latro, or represent a shift in emphasis of higher order neuropils and processing capability toward olfactory instead of tactile inputs.

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David C. Sandeman, Matthes Kenning, and Steffen Harzsch Parasol cells Type 1 Type 2 PC

Ch em o M ech senso HBN1 Vis anos ry Ch ual ens TM ory em ose nso ry HBN2

Visual Mechanosensory

Local interneurons LNrim (9) LNcore

ORN Chemosensory

Cap Core

Contralaterals

OL glomeruli

GN

(11) Base

OGT

(10) Projection neurons

DC

AL

Mechanosensory, visual

Medulla Cortex

Fig. 2.6. Summary of the known connections between the lateral deutocerebral and lateral protocerebral neuropils of crayfish and clawed lobsters. The olfactory receptor neurons are the only primary sensory afferent input, and these end in the glomeruli of the olfactory lobe. The accessory lobe, terminal medulla, and hemiellipsoid body receive chemical, mechanosensory, and visual input via interneurons. Olfactory information is relayed from the olfactory lobe to the cortex of the accessory lobe, to the terminal medulla, and to the outer neuropil of the hemiellipsoid body. Visual information reaches the medulla of the accessory lobe and the inner neuropil of the hemiellipsoid body. Mechanosensory information is directed to the medulla of the accessory lobe, to the terminal medulla, and to the inner neuropil of the hemiellipsoid body. Identified local interneurons are associated with the olfactory and accessory lobes (local olfactory interneurons, dorsal giant neuron, and midbrain olfactory interneurons) and with the hemiellipsoid bodies (parasol cells). (Based on the results of Derby and Blaustein 1988, Blaustein et al. 1988, Sandeman and Sandeman 1994, Mellon and Alones 1995, 1997, Sandeman D. et al. 1995, Sullivan and Beltz 2001, 2005, McKinzie et al. 2003, Mellon 2007). Abbreviations: AL, accessory lobe; DC, deutocerebrum; GN, dorsal giant neuron; HBN12, subdivision of the hemiellipsoid body neuropil; LN, local interneuron; OGT, olfactory globular tract; ORN, olfactory receptor neuron; PC, protocerebrum; TM, terminal medulla.

Accessory Lobes The spiny lobsters have large and complex accessory lobes. These lie medial to the olfactory lobes and have three layers of glomerular neuropil (Blaustein et al. 1988) in which those in the outer layer are columnar, while those in the inner two layers are spherical (Sandeman et al. 1992,



Adaptive Trends in Malacostracan Brain Form and Function Related to Behavior

Wachowiak et al. 1996, Schmidt and Ache 1997). The accessory lobes of the astacids have two concentric layers of glomerular neuropil that are all spherical in crayfish (Sandeman D. et al. 1995). In the clawed lobster H. americanus, the glomeruli in the outer layer are columnar like those found in the spiny lobsters, and spherical in the inner layer (Helluy et al. 1993). The accessory lobes receive no primary afferent inputs, and the input from the olfactory lobes, via local interneurons, terminates predominantly in the cortical areas (Sullivan and Beltz 2005). The accessory lobes in C.  destructor also receive inputs from a large number of interneurons in the deutocerebral commissure. Intracellular electrical recordings and subsequent labeling of these interneurons with neurobiotin revealed five anatomical classes of interneurons. These have unilateral inputs located in different areas in the proto- or deutocerebrum or bilateral inputs from the tritocerebrum, and all have bilateral outputs ending in the glomeruli of the left and right accessory lobes. Information from olfactory, visual, and tactile inputs are represented in the axons of the interneurons of the deutocerebral commissure (Sandeman D. et al. 1995), and ultrastructural studies of labeled deutocerebral commissure interneurons confirmed them to be presynaptic to elements within the accessory lobe (Sandeman R. et al. 1995). The accessory lobes in B. latro on the other hand, although containing small spherical glomeruli like those in the crayfish, are very small and do not appear to be connected to each other or to other areas of the brain via a deutocerebral commissure. Given their very small volume, it is difficult to see how they could play a dominant role in the olfactory processing in these animals such as that proposed for the crayfish or lobsters. The emphasis on the olfactory lobes and the hemiellipsoid bodies in B. latro, and the extreme reduction of the accessory lobes, suggest a shift in the balance of processing power. Spiny lobsters, and some crayfish with equally long second antennae, have significantly larger accessory lobes compared with those species relying on olfactory and visual senses. Hence the accessory lobes could be integrative areas also serving exploratory mechano-chemosensory behavior (see Conclusions). Evolution of the Malacostracan Higher Integrative Centers Early carcinologists identified the olfactory lobes, accessory lobes, and lateral protocerebrum as areas where considerable evolutionary transformation has taken place during phylogeny of Malacostraca. Olfactory lobes were most likely present in the ground pattern of Malacostraca and linked to the lateral protocerebrum by the protocerebral tract (Fig. 2.2A). In the Decapoda, the projection neurons probably equally innervated both the terminal medullae and the hemiellipsoid bodies, and contralateral projections of the olfactory globular tract were present as in the original malacostracan ground pattern (Fig. 2.7). This configuration could be homologous to that of basal insects, which lack mushroom bodies (Farris 2005, Galizia and Rössler 2010). Their antennal lobes (homologs of the malacostracan olfactory lobes) are linked to the lateral horn in the protocerebrum (a homolog of the malacostracan lateral protocerebrum), but the antennocerebral tract is uncrossed (Galizia and Rössler 2010). During evolution of the Pterygota, the functional need for more sophisticated olfactory processing arose to cope with airborne odors, and insects developed mushroom bodies (only the peduncles in the beginning) to provide an adequate neuronal substrate. They also evolved additional projection neuron tracts that link the antennal lobes to the secondary olfactory areas in the protocerebrum (Fig. 2.7; Galizia and Rössler 2010, Strausfeld 2012). The Malacostraca appear to have followed a different pathway to increase their capacity for analyzing olfactory stimuli. In the Reptantia, the accessory lobe, an evolutionary novelty, arose. Most of the projection neurons associated with the accessory lobes in Reptantia target the hemiellipsoid bodies, whereas those from the olfactory lobe are connected predominantly to the terminal medullae (Sullivan and Beltz 2001, 2005).

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David C. Sandeman, Matthes Kenning, and Steffen Harzsch Crustacea

Hexapoda

Malacostraca HBN PC TM

Decapoda

iAcT

LH Reptantia

OL

Antennal lobe

MB

DC

oAcT AL

Coenobitidae

Brachyura

Astacida HBN2 HBN1

Homarida HBN3 HBN2 HBN1

}

Medulla AL Cortex

Fig. 2.7. Hypothesis on the evolution of the malacostracan and insect higher integrative centers (for further details see text). Abbreviations:  AL, accessory lobe; DC, deutocerebrum; HBN, hemiellipsoid body; HBN1-3, subdivision of the hemiellipsoid body neuropil; iAcT, inner antennocerebral tract; LH, lateral horn; MB, mushroom body; oAcT, outer antennocerebral tract; OL, olfactory lobe; PC, protocerebrum; TM, terminal medulla. Question mark denotes uncertainty about the connection.

The accessory lobes first appeared in early Reptantia, then lost their presumed role as secondary olfactory centers in Brachyura and Anomura. The development and then loss of the accessory lobes is difficult to resolve in terms of lifestyle and habitat although there are differences between the groups that may provide some clues. Many of the Achelata, Homarida, and Astacida are either nocturnal or crepuscular and have well developed second antennae and good mechanosensory abilities. Many of the Brachyura on the other hand have well-developed visual systems but the second antennae are often relatively short. Many are active both at night and during the day. In terms of the proportional sensory modalities, the animals would appear to rely on a combination of olfactory and visual inputs. The Thalassinida represents an intermediate stage: Callianassa australiensis has enlarged hemiellipsoid bodies that are located medially in the brain and accessory lobes that, while smaller than those of the Astacida, are linked to one another across the brain by a well-developed deutocerebral commissure (Sandeman and Scholtz 1995).



Adaptive Trends in Malacostracan Brain Form and Function Related to Behavior

The Coenobitidae in this context is an interesting case because this group of terrestrial hermit crabs, which includes the genus Coenobita and the monospecific genus Birgus, has secondarily emphasized the olfactory pathway (Harzsch and Hansson 2008, Krieger et al. 2010). There is good behavioral and physiological evidence for a sense of aerial olfaction in members of the Coenobitidae (review Hansson et al. 2011), and their olfactory lobes are large. The hemiellipsoid bodies are also characteristically layered and large in comparison with those found in most other decapods (Fig. 2.2). In summary, it appears that in Insecta and Malacostraca, we find three different strategies to increase the analytical and integrative need to match the shifting proportion of sensory inputs (Fig. 2.7). This could be shared between visual, olfactory, and mechanosensory or dominated by any one, or a pair of inputs. In terms of the chemosensory input: (1) introduce a new neuropil in the protocerebrum and hook it up to the central olfactory pathway (Pterygota), (2) introduce a new neuropil in the deutocerebrum (Reptantia) and hook it up to the central olfactory pathway, and (3) inflate the existing neuronal substrate of the central olfactory pathway and elaborate its architecture (Coenobitidae).

FUTURE DIRECTIONS Classical methods have provided us with a basic morphological and physiological framework in which to understand the brains and behavior of some of the larger decapod crustaceans. Advances beyond this depend on the development and application of new methods and techniques of which there are already an exciting range from which to choose. For example, miniaturization of video and radio tracking devices as well as GPS-based telemetry systems now allow ethologists to move into the field and beyond the observation of animals confined in enclosures or aquaria. Developments in immunocytochemistry provide chemical snapshots of the ebb and flow of a host of neurochemical agents that can be related to behavioral activities. The introduction of laser scanning microscopes and fluorescent dyes has considerably extended the resolution of conventional microscopes. Three-dimensional reconstructions of image stacks either obtained from confocal microscopy or serial sections add significant new possibilities to neuroanatomical analysis, enabling comparative and evolutionary neuroanatomists to rapidly screen and explore morphological variations in many different species. Recent advances in computer tomography permit the relatively noninvasive observation of brain chemistry in real time, thus freeing physiologists from the need to impale living tissues and interpret the neuronal chemistry from associated electrical signals. Such innovative approaches may be expected to advance our understanding in terms of the phylogenetic relationships within the Crustacea as a whole and also of the roles played by the “difficult” areas of the crustacean brains, such as the hemiellipsoid bodies and the accessory and olfactory lobes. These, by virtue of their sheer complexity and large numbers of small neurons contained within them, have resisted the attempts of physiologists to unravel their connectivity or provide a coherent explanation of their functional roles in crustacean behavior. It is here that the application of the new wave of technologies could have the greatest impact.

CONCLUSIONS Brain and Behavior in the Malacostracans In comparing the neuropils within the Malacostraca, one feature seems to be clear: the high level of conservation of the characteristic anatomical features that we use to recognize the

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David C. Sandeman, Matthes Kenning, and Steffen Harzsch neuropils. This may be explained to some extent by a consideration of the roles of these neuropils and the type of modality that processing within them extracts from the receptor cells that project to them. In visual systems, the information from an array of photoreceptors that can merely respond to light intensity is translated into the important properties that constitute vision, namely extraction from the photoreceptor array of the contrasting boundaries that go to make up objects and the interpretation of the direction and velocity of the movement of this object. To achieve this, the information from the photoreceptors passes through layers of elements where specific lateral interactions occur and where the nature of the information that is extracted gradually becomes more sophisticated, changing from mere light intensity to the detection of differences between areas, the sensitivity only to edges or lines in particular directions, and the direction and velocity of motion. This is the case in all visual systems, both vertebrate and nonvertebrate. Hence, it is perhaps no surprise to find that despite the vast difference in the visually guided behavior patterns of stomatopods and swimming crabs on the one hand and that of the isopods on the other, the actual anatomical characteristics of the layered optic neuropil appear very similar. The essential differences in terms of the abilities of the two groups presumably lie in the numbers and in the tuned responses of the individual neurons that are contained in the various layers. The differences between the animals’ visual systems are therefore not qualitative (they are all capable of detecting the direction and velocity of a moving edge) but quantitative in terms of their precision. This argument also applies to the olfactory systems. Olfactory receptor cells in many animals are supplied with unique molecular receptors that constitute the first filter in the system in selecting specific odors, much like photoreceptors are able to selectively respond to different wavelengths of light. There is a precise projection of these cells, as in the visual system, to discrete areas of the olfactory lobe neuropil where specific processing will extract the chemical information that is meaningful for the animal. As in the case of the visual system, the structure of malacostracan olfactory systems is highly conserved, consisting of tight areas of neuropil or glomeruli. From these areas that receive the primary afferent inputs, there are trunks of large numbers of fine axons, the projection neurons, that extend to a second area of neuropil in the protocerebrum, the hemiellipsoid bodies, which in some species (B. latro, for example) have layers of laterally projecting neurons. The extraction of the relevant information in exploratory mechano-chemosensory systems may be equally complex in comparison with vision and olfaction. The antenna 2 neuropil in animals with long flexible second antennae is geometrically structured and may provide the substrate for the spatiotopic location of antennal stimulation. If animals are employing their second antennae, and other appendages (legs) for the active mechano-chemosensory exploration of their surroundings, this would require a high degree of central integration incorporating not only the comparison of motor commands with the ensuing sensory responses, but also including visual and olfactory inputs. Where is the relevant higher order neuropil for this system? Perhaps the assignment of the accessory lobes purely to the olfactory pathway is an oversimplification and its real function extends to the reception of the wide range of information gathered during the active examination of objects and the surroundings, the evaluation of this in relation to previous experience and the generation of appropriate action. Functional connections between the antenna 2 neuropils, the visual system, the olfactory lobes, and the accessory lobes have been described in C. destructor (Sandeman D. et al. 1995) and are highly likely to be also present in other species with accessory lobes. Support for a highly integrative and behaviorally relevant role for the accessory lobes has come from a recent exploration of drug dependency which used crayfish as a model system: an



Adaptive Trends in Malacostracan Brain Form and Function Related to Behavior

alteration of the c-Fos mRNA expression in the accessory lobe of crayfish has been associated with a conditioned cocaine-induced reward. The remarkable aspect of this study in the context of the function of the accessory lobes is that textural information from the substratum on which the animals were placed was coupled with the cocaine conditioning. This suggests that the mechanoreceptive differences in the substratum (pebbled or smooth) detected by the walking legs are relayed to the accessory lobes, which in turn are implicated in the behavioral changes associated with the drug administration (Nathaniel et al. 2012). Further evidence for the multifunctionality of the accessory lobes comes from the discovery of a neurogenic niche associated with the accessory lobes of many of the reptantians (Bazin 1970, Sandeman et al. 2011, c­ hapter 7 in this volume). Best described in the crayfish P. clarkii (Benton et  al. 2011, Beltz et  al. 2011)  and spiny lobster P.  argus (Schmidt 2007, Schmidt and Derby 2011), this system generates new local and projection neurons throughout the life of the adult animals, and these new neurons are incorporated into the olfactory pathway (see ­chapter 7 in this volume). Multimodality, Convergence, and Cognition Spiny lobsters and coconut crabs are both large and long-lived species of crustaceans, and both exhibit behaviors that are complex and of long duration and in which a particular sequence is essential in order for the ultimate behavioral aim to be achieved. Migrations must be initiated and directed, intermediate localities and appropriate releasers recognized and acted on, and finally home journeys initiated, undertaken, and home localities found. Behaviors of this kind, with the unpredictable intermediate events that could occur during the completion of the task, cannot be undertaken without some level of learning and recognition of a particular constellation of olfactory, visual, tactile, and geomagnetic stimuli. The actual task of combining these features into a unique set that can be matched with some preexisting “image” is no different for a homing spiny lobster or crab than that confronting any “higher” organism equipped with a cortex. To combine a set of various sensory stimuli into some unique neuronal picture requires the presence of multimodal neurons that will respond to a particular set of inputs that may occur in a particular sequence and at particular levels of intensity. Certainly there are many examples of these kinds of neurons at all levels in the brains of the crustaceans and particularly in the accessory lobe, hemiellipsoid body, and terminal medulla. These may be responsible for a highly selective filtering of sets of environmental inputs and trigger an appropriate behavior. In other words, simple convergence, given adequate and perhaps tuned preset thresholds, could do the job, and the brain areas containing these are represented in the crustacean central nervous system (Fig. 2.6). There are, however, some difficulties with such a simple model. In a consideration of how multiple sensory inputs are integrated in the mammalian cortex, Tonini et al. (1998) point out that convergence is unlikely to be the predominant mechanism because no “master” brain area has been identified and there are so many possible combinations of stimuli, each of which would need its own master area, that there are not even enough single neurons available to cover the combinatorial explosion that would ensue (and certainly not in the relatively small brains of the arthropods). The same authors point out that convergence will not allow the flexibility needed to respond to novel stimuli, and in the context of a coconut crab finding its way to the coast to mate, or a spiny lobster migrating offshore, it is certain that the detailed features of the landscape through which they make their way are not going to be precisely the same in subsequent migrations. Instead of simple convergence of multimodal inputs, it seems that a more likely solution would include interactions between functionally segregated brain areas that act to synchronize

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David C. Sandeman, Matthes Kenning, and Steffen Harzsch the ongoing activity of groups of neurons (Tonini et al. 1998). A feature of such models is that target cells are not driven by such connections but that the timing of their firing is modulated. In this regard, the parasol cells in the hemiellipsoid body of the crayfish provide an interesting parallel (Mellon and Wheeler 1999, Mellon 2000, McKinzie et al. 2003). These neurons fire spontaneously, exhibit particular and individual patterns, and, most important for the comparison with the cortex model, these patterns of activity are modulated rather than directly driven by their multimodal receptor inputs. They could therefore represent the morphological substrate for the “reentrant” requirement of the cortex model, in which parallel and collateral interactions between separate brain areas occur. Furthermore, the cortex model, which satisfies many experimental findings, contains elements such as “functional clusters” that are defined as a “set of brain regions that interact much more strongly with each other than with the rest of the brain” (Tonini et al. 1998). The accessory lobe, the hemiellipsoid body, and terminal medulla in the malacostracans are candidates for such functions and worth exploring in this context.

REFERENCES Alwes, F., and G. Scholtz. 2006. Stages and other aspects of the embryology of the parthenogenetic Marmorkrebs (Decapoda, Reptantia, Astacida). Development Genes and Evolution 216:169–184. Atema, J., and J.S. Cobb. 1980. Social behavior. Pages 409–450 in J.S. Cobb and B.F. Phillips, editors. The biology and management of lobsters, Vol. 1. Academic Press, New York. Basil J., and D. Sandeman. 2000. Crayfish (Cherax destructor) use tactile cues to detect and learn topographical changes in their environment. Ethology 106:247–259. Bazin, F. 1970. Étude comparée de ľorgane deutocérébrale des Macroures Reptantia et des Anomoures (Crustacés Décapodes). Archives de Zoologie Expérimentale et Générale 11:245–264. Becker, J.H.A., L.M. Curtis, and A.S. Grutter. 2005. Cleaner shrimps use a rocking dance to advertise cleaning service to clients. Current Biology 15:760–764. Beltz, B.S. 1999. The distribution and functional anatomy of amine-containing neurons in decapod crustaceans. Microscopy Research and Technique 44:105–120. Beltz, B.S., K. Kordas, M.M. Lee, J.B. Long, J.L. Benton, and D.C. Sandeman. 2003. Ecological, evolutionary and functional correlates of sensilla number and glomerular density in the olfactory system of decapod crustaceans. Journal of Comparative Neurology 455:260–269. Beltz, B.S., Y. Zhang, J.L. Benton, and D.C. Sandeman. 2011. Adult neurogenesis in the decapod crustacean brain: a hematopoietic connection? European Journal of Neuroscience 34:670–883. Bender, M., W. Gnatzy, and J. Tautz. 1984. The antennal feathered hairs in the crayfish: a non-innervated stimulus transmitting system. Journal of Comparative Physiology A 154:45–47. Benton, J.L., Y. Zhang, C.R. Kirkhart, D.C. Sandeman, and B.S. Beltz. 2011. Primary neuronal precursors in adult crayfish brain: replenishment from a non-neuronal source. BMC Neuroscience 12:53 Berry, F.C., and T. Breithaupt. 2010. To signal or not to signal? Chemical communication by urine-borne signals mirrors sexual conflict in crayfish. BMC Biology 8:25. Blaustein, D.N., C.D. Derby, R.B. Simmons, and A.C. Beall. 1988. Structure of the brain and medulla terminalis of the spiny lobster Panulirus argus and the crayfish Procambarus clarkii, with an emphasis on olfactory centers. Journal of Crustacean Biology 8:493–519. Boles L.C., and K.J. Lohmann. 2003. True navigation and magnetic maps in spiny lobsters. Nature 421:60–63. Breithaupt, T. 2001. Fan organs of crayfish enhance chemical information flow. Biological Bulletin 200:150–154. Breithaupt, T. 2011. Chemical communication in crayfish. Pages 257–276 in T. Breithaupt and M. Thiel, editors. Chemical communication in crustaceans. Springer, New York. Breithaupt, T., and M. Thiel, editors. 2011. Chemical communication in crustaceans. Springer. New York.



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Briones-Fourzán, P., M. Pérez-Ortiz, and E. Lozano-A lvarez. 2006. Defense mechanisms and antipredator behavior in two sympatric species of spiny lobster, Panulirus argus and P. guttatus. Marine Biology 149:227–239. Briones-Fourzán, P., E. Ramírez-Zaldívar, and E. Lozano-A lvarez. 2008. Influence of conspecific and heterospecific aggregation cues and alarm odors on shelter choice by syntopic spiny lobsters. Biological Bulletin 215:182–190. Burrows, M., and G.A. Horridge. 1968. The action of the eyecup muscles of the crab Carcinus during optokinetic movements. Journal of Experimental Biology 49:223–250. Bushmann, P.J., and J. Atema. 1996. Nephropore rosette glands of the lobster Homarus americanus: possible sources of urine pheromones. Journal of Crustacean Biology 16:221–231. Caldwell, R.L., and H. Dingle. 1975. Ecology and evolution of agonistic behavior in stomatopods. Naturwissenschaften 62:214–222. Cate, H.S., and C.D. Derby. 2002. Hooded sensilla homologues: structural variations of a widely distributed bimodal chemo-mechanosensillum. Journal of Comparative Neurology 444:345–357. Childress, M.J., and W.F. Herrnkind. 2001. The guide effect influence on the gregariousness of juvenile Caribbean spiny lobsters. Animal Behaviour 62:465–472. Cooper, R.L., H. Li, L.Y. Long, J.L. Cole, and H. Hopper. 2001. Anatomical comparisons of neural systems in sighted epigean and troglobytic crayfish species. Journal of Crustacean Biology 21:360–374. Cox, C., J.H. Hunt, W.G. Lyons, and G.E. Davis. 1997. Nocturnal foraging of the Caribbean spiny lobster (Panulirus argus) on offshore reefs of Florida, USA. Marine and Freshwater Research 48:671–679. Creaser, E.P., and D. Travis. 1950. Evidence of homing instinct in the Bermuda spiny lobster. Science 112:169–170. Cronin, T.W., J.N. Nair, R.D. Doyle, and R.L. Caldwell. 1988. Ocular tracking of rapidly moving visual targets by stomatopod crustaceans. Journal of Experimental Biology 138:155–179. Derby, C.D., and D.N. Blaustein. 1988. Morphological and physiological characterization of individual olfactory interneurons connecting the brain and eyestalk ganglia of the crayfish. Journal of Comparative Physiology A 163:777–794. Derby, C.D., H.S. Cate, and L.R. Gentilcore. 1997. Perireception in olfaction: molecular mass sieving by aesthetasc sensillar cuticle determines odorant access to receptor sites in the Caribbean spiny lobster Panulirus argus. Journal of Experimental Biology 200:2073–2081. Drew, M.M., S. Harzsch, M. Stensmyr, S. Erland, and B.S. Hansson. 2010. A review of the biology and ecology of the Robber Crab, Birgus latro (Linnaeus, 1767) (Anomura: Coenobitidae). Zoologischer Anzeiger 249:45–67. Farris, S.M. 2005. Evolution of insect mushroom bodies: old clues, new insights. Arthropod Structure and Development 34:211–234. Galizia, C.G., and W. Rössler. 2010. Parallel olfactory systems in insects: anatomy and function. Annual Review of Entomology 55:399–420. Grutter, A.S. 2004. Cleaner fish use tactile dancing behavior as a preconflict management strategy. Current Biology 14:1080–1083. Hallberg, E., and M. Skog. 2011. Chemosensory sensilla in crustaceans. Pages 103–121 in T. Breithaupt and M. Thiel, editors. Chemical communication in crustaceans. Springer, New York. Hallberg, E., K.U.I. Johansson, and R. Elofsson. 1992. The aesthetasc concept: structural variations of putative olfactory receptor cell complexes in Crustacea. Microscopy Research and Technique 22:325–335. Hansson, B.S., S. Harzsch, M. Knaden, and M. Stensmyr. 2011. The neural and behavioral basis of chemical communication in terrestrial crustaceans. Pages 149–173 in T. Breithaupt and M. Thiel, editors. Chemical communication in crustaceans. Springer, New York. Harzsch, S., and B.S. Hansson. 2008. Brain architecture in the terrestrial hermit crab Coenobita clypeatus (Anomura, Coenobitidae), a crustacean with a good aerial sense of smell. BMC Neuroscience 9:58. Harzsch, S., V. Rieger, J. Krieger, F. Seefluth, N.J. Strausfeld, and B.S. Hansson. 2011. Transition from marine to terrestrial ecologies: changes in olfactory and tritocerebral neuropils in land-living isopods. Arthropod Structure and Development 40:251–270.

43

44

David C. Sandeman, Matthes Kenning, and Steffen Harzsch Hay, M.E. 2011. Crustaceans as powerful models in aquatic chemical ecology. Pages 41–62 in T. Breithaupt and M. Thiel, editors. Chemical communication in crustaceans. Springer, New York. Helluy, S., R. Sandeman, B. Beltz, and D. Sandeman. 1993. Comparative brain ontogeny of the crayfish and clawed lobster: implications of direct and larval development. Journal of Comparative Neurology 335:343–354. Herrnkind, W.F., and R. McLean. 1971. Field studies of orientation, homing, and mass migration in the spiny lobster, Panulirus argus. Annals of the New York Academy of Sciences 188:359–377. Herrnkind, W.F., J.A. Vanderwalker, and L. Barr. 1975. Population dynamics, ecology and behavior of spiny lobsters, Panulirus argus, of St. John, USVI. IV. Habitation, patterns of movement and general behavior. Results of Tektite Program. Natural History Museum of Los Angeles County Science Bulletin 20:31–45. Herrnkind, W.F., M.J. Childress, and K.L. Lavalli. 2001. Cooperative defence and other benefits among exposed spiny lobsters: inferences from group size and behaviour. Marine and Freshwater Research 52:1113–1124. Horner, A.J., M.J. Weissburg, and C.D. Derby. 2008. The olfactory pathway mediates sheltering behavior of Caribbean spiny lobsters (Panulirus argus). Journal of Comparative Physiology A 194:243–253. Horridge, G.A. 1978. The separation of visual axes in apposition compound eyes. Philosophical Transactions of the Royal Society of London B 285:1–59. Jernakoff, P. 1987. Foraging patterns of juvenile western rock lobsters Panulirus cygnus George. Journal of Experimental Marine Biology and Ecology 113:125–144. Jones, J. 1994. Architecture and composition of the muscles that drive stomatopod eye movements. Journal of Experimental Biology 188:317–331. Katsanevakis, S., and G. Verriopoulos. 2006. Seasonal population dynamics of Octopus vulgaris in the eastern Mediterranean. ICES Journal of Marine Science 63:151–160. Kleinlogel, S., N.J. Marshall, L.M. Horwood, and M.F. Land. 2003. Neuroarchitecture of the color and polarization vision system of the stomatopod Haptosquilla. Journal of Comparative Neurology 467:326–342. Krieger, J., R.E. Sandeman, D.C. Sandeman, B.S. Hansson, and S. Harzsch. 2010. Brain architecture of the largest living land arthropod, the giant robber crab Birgus latro (Crustacea, Anomura, Coenobitidae): evidence for a prominent central olfactory pathway? Frontiers in Zoology 7:25. Krieger, J., A. Sombke, F. Seefluth, M. Kenning, B.S. Hansson, and S. Harzsch. 2012. Comparative brain architecture of the European shore crab Carcinus maenas (Brachyura) and the common hermit crab Pagurus bernhardus (Anomura). Cell and Tissue Research 384:47–69. Land, M.F., N.J. Marshall, D. Brownless, and T.W. Cronin. 1990. The eye movements of the mantis shrimp Odontodactylus scyllarus (Crustacea: Stomatopoda). Journal of Comparative Physiology A 167:155–166. Larimer, J.L. 1966. A functional caudal photoreceptor in blind caverniculous crayfish. Nature 21:204–205. Li, H., and R.L. Cooper. 1999. The neuroecology of the blind cave crayfish: social interactions. American Zoologist 38:201A. Li, H., L.R. Listeman, D. Doshi, and R.L. Cooper. 2000. Heart rate measures in blind crayfish during environmental and social interactions. Comparative Biochemistry and Physiology 127A:55–70. Linsenmaier, K.E. 1987. Kin recognition in subsocial arthropods, in particular in the desert isopod Hemilepistus reamuri. Pages 121–207 in D. Fletcher and C. Michener, editors. Kin recognition in animals. John Wiley and Sons, Chichester. Linsenmaier, K.E. 2007. Sociobiology of terrestrial isopods. Pages 339–364 in J.E. Duffy and M. Thiel, editors. Evolutionary ecology of social and sexual systems: crustaceans as model organisms. Oxford University Press, Oxford. Lohmann, K.J., N.D. Pentcheff, G.A. Nevitt, G.D. Stetten, R.K. Zimmer-Faust, H.E. Jarrard, and L.C. Boles. 1995. Magnetic orientation of spiny lobsters in the ocean: experiments with undersea coil systems. Journal of Experimental Biology 198:2041–2048. Marshall, N.J., J.P. Jones, and T.W. Cronin. 1996. Behavioural evidence for colour vision in stomatopod crustaceans. Journal of Comparative Physiology A 179:473–481.



Adaptive Trends in Malacostracan Brain Form and Function Related to Behavior

Marshall, J., T.W. Cronin, N. Shashar, and M. Land. 1999. Behavioural evidence for polarisation vision in stomatopods reveals a potential channel for communication. Current Biology 9:755–758. Marshall, J., T.W. Cronin, and S. Kleinlogel. 2007. Stomatopod eye structure and function: a review. Arthropod Structure and Development 36:420–448. Masters, W.M., B. Aicher, J. Tautz, and H. Markl. 1982. A new type of water vibration receptor on the crayfish antenna. II. Model of receptor function. Journal of Comparative Physiology A 149:409–422. McKinzie, M.E., J.L. Benton, B.S. Beltz, and DeF. Mellon. 2003. Parasol cells of the hemiellipsoid body in the crayfish Procambarus clarkii: dendritic branching patterns and functional implications. Journal of Comparative Neurology 462:168–179. McMahon, A., B.W. Patullo, and D.L. Macmillan. 2005. Exploration in a T-maze by the crayfish Cherax destructor suggests bilateral comparison of antennal tactile information. Biological Bulletin 208:183–188. Mead, K., and R. Caldwell. 2011. Mantis shrimp: olfactory apparatus. Pages 219–238 in T. Breithaupt and M. Thiel, editors. Chemical communication in crustaceans. Springer, New York. Mellon, DeF. Jr. 1977. Retention of oculomotor reflexes in blind cave-dwelling crayfish. Brain Research 134:191–196. Mellon, DeF. Jr. 2000. Convergence of multimodal sensory input onto higher-level neurons of the crayfish olfactory pathway. Journal of Neurophysiology 84:3043–3055. Mellon, DeF. Jr. 2003. Active dendrite properties constrain input-output relationships in neurons of the central olfactory pathway in the crayfish forebrain. Microscopy Research and Technique 60:278–290. Mellon, DeF. Jr. 2007. Combining dissimilar senses: central processing of hydrodynamic and chemosensory inputs in aquatic crustaceans. Biological Bulletin 213:1–11. Mellon, DeF. Jr., and V. Alones. 1993. Cellular organization and growth-related plasticity of the crayfish olfactory midbrain. Microscopy Research and Technique 24:231–259. Mellon, DeF. Jr., and V. Alones. 1995. Identification of three classes of multiglomerular, broad spectrum neurons in the crayfish olfactory midbrain by correlated patterns of electrical activity and dendritic arborization. Journal of Comparative Physiology A 177:55–71. Mellon, DeF. Jr., and V. Alones. 1997. Response properties of higher-level neurons in the central olfactory pathway of the crayfish. Journal of Comparative Physiology A 181:205–216. Mellon, DeF. Jr., and G. Lnenicka. 1980. Structure and electrical properties of eye muscles in cave- and surface-dwelling crayfishes. Journal of Experimental Biology 84:187–199. Mellon, DeF. Jr., and M.A. Reidenbach. 2011. Fluid mechanical problems in crustacean active chemoreception. Pages 159–170 in Frontiers in sensing: from biology to engineering. F. Barth, J.A.C. Humphrey, and M.V. Srinivasan, editors. Springer Verlag, Wien, New York. Mellon, DeF. Jr., and C.J. Wheeler. 1999. Coherent oscillations in membrane potentials synchronize impulse bursts in central olfactory neurons of the crayfish. Journal of Neurophysiology 81:1231–1241. Mellon, DeF. Jr., V. Alones, and M.D. Lawrence. 1992a. Anatomy and fine structure of neurons in the deutocerebral projection pathway of the crayfish olfactory system. Journal of Comparative Neurology 321:93–111. Mellon, DeF. Jr., D.C. Sandeman, and R.E. Sandeman. 1992b. Characterization of oscillatory olfactory interneurones in the protocerebrum of the crayfish. Journal of Experimental Biology 167:15–38. Moore, P.A. 2007. Agonistic behavior in freshwater crayfish: the influence of intrinsic and extrinsic factors on aggressive encounters and dominance. Pages 90–11 in J.E. Duffy and M. Thiel, editors. Evolutionary ecology of social and sexual systems: crustaceans as model organisms. Oxford University Press, Oxford. Nathaniel, T.I., J. Panksepp, and R. Huber. 2012. Alteration of c-Fos mRNA in the accessory lobe of crayfish is associated with a conditioned-cocaine induced reward. Neuroscience Research 72:243–256. Olszewski, P. 1980. A salute to the humble yabby. Angus and Robertson, Sydney. Page, T.L. 1982. Extraretinal photoreception in entrainment and photoperiodism in invertebrates. Experientia 38:1007–1013.

45

46

David C. Sandeman, Matthes Kenning, and Steffen Harzsch Peters, W.L., and I.C. Campbell. 1991. Ephemeroptera (mayflies). Pages 279–293 in I.D. Naumann, editor. The insects of Australia, Vol.1. Melbourne University Press, Melbourne. Reynolds, K.M. 1980. Aspects of the biology of the freshwater crayfish, Cherax destructor in farm dams in far-western NSW. Master of Science Thesis. University of New South Wales, NSW, Australia. Richter, S., and G. Scholtz. 2001. Phylogenetic analysis of the Malacostraca (Crustacea). Journal of Zoological Systematics and Evolutionary Research 39:113–136. Rossi-Durand, C., and J.P. Vedel. 1982. Antennal proprioception in the rock lobster Palinurus vulgaris: anatomy and physiology of a bi-articular chordotonal organ. Journal of Comparative Physiology A 145:505–516. Sandeman, D.C. 1985. Crayfish antennae as tactile organs: their mobility and the response of their proprioceptors to displacement. Journal of Comparative Physiology A 157:363–373. Sandeman, D.C. 1989. Physical properties, sensory receptors, and tactile reflexes of the antenna of the crayfish Cherax destructor. Journal of Experimental Biology 141:197–217. Sandeman, D.C., and J.L. Denburg. 1976. The central projections of the chemoreceptor axons in the crayfish revealed by axoplasmic transport. Brain Research 115:492–496. Sandeman, D.C., and S.E. Luff. 1973. The structural organization of glomerular neuropile in the olfactory and accessory lobes of an Australian freshwater crayfish. Zeitschrift für Zellforschung und Mikroskopische Anatomie 142:37–61. Sandeman, D.C., and R.E. Sandeman. 1994. Electrical responses and synaptic connections of giant serotonin-immunoreactive neurons in crayfish olfactory and accessory lobes. Journal of Comparative Neurology 342:130–144. Sandeman, D.C., and G. Scholtz. 1995. Ground plans, evolutionary changes and homologies in Decapod crustacean brains. Pages 329–348 in W. Kutsch and O. Breidbach, editors. The nervous systems of invertebrates: a comparative approach. Birkhäuser, Basel. Sandeman, D.C., and D. Varju. 1988. A behavioural study of tactile localization in the crayfish Cherax destructor. Journal of Comparative Physiology A 163:525–536. Sandeman D.C., R.E. Sandeman, and H.G. de Couet. 1990. Extraretinal photoreceptors in the brain of the crayfish Cherax destructor. Journal of Neurobiology 21:619–629. Sandeman, D.C., R.E. Sandeman, C. Derby, and M. Schmidt. 1992. Morphology of the brain of crayfish, crabs and spiny lobsters: a common nomenclature for homologous structures. Biological Bulletin 183:304–326. Sandeman, D.C., G. Scholtz, and R.E. Sandeman. 1993. Brain evolution in decapod crustaceans. Journal of Experimental Zoology 265:112–133. Sandeman, D., B. Beltz, and R. Sandeman. 1995. Crayfish brain interneurons that converge with serotonin giant cells in the accessory lobe glomeruli. Journal of Comparative Neurology 352:263–279. Sandeman, D.C., F. Bazin, and B.S. Beltz. 2011. Adult neurogenesis: examples from the decapod crustaceans and comparisons with mammals. Arthropod Structure and Development 40:258–275. Sandeman, R.E., and D.C. Sandeman. 1996. Pre- and postembryonic development, growth and turnover of olfactory receptor neurons in crayfish antennules. Journal of Experimental Biology 199:2409–2418. Sandeman, R.E., A.H.D. Watson, and D.C. Sandeman. 1995. Ultrastructure of the synaptic terminals of the dorsal giant serotonin-IR neuron and deutocerebral commissure interneurons in the accessory and olfactory lobes of the crayfish. Journal of Comparative Neurology 361:617–632. Schachtner, J., M. Schmidt, and U. Homberg. 2005. Organization and evolutionary trends of primary olfactory brain centers in Tetraconata (Crustacea + Hexapoda). Arthropod Structure and Development 34:257–299. Schaller, F. 1953. Verhaltens- und sinnesphysiologische Beobachtungen an Squilla mantis. Zeitschrift für Tierpsychologie 10:1–12. Schmidt, M. 2007. The olfactory pathway of decapod crustaceans—an invertebrate model for life-long neurogenesis. Chemical Senses 32:365–384. Schmidt, M., and B.W. Ache. 1997. Immunocytochemical analysis of glomerular regionalization and neuronal diversity in the olfactory deutocerebrum of the spiny lobster. Cell and Tissue Research 287:541–563.



Adaptive Trends in Malacostracan Brain Form and Function Related to Behavior

Schmidt, M., and C.D. Derby. 2011. Cytoarchitecture and ultrastructure of neural stem cell niches and neurogenic complexes maintaining adult neurogenesis in the olfactory midbrain of spiny lobsters, Panulirus argus. Journal of Comparative Neurology 519:2283–2319. Schmidt, M., and DeF. Mellon, Jr. 2011. Neuronal processing of chemical information in crustaceans. Pages 123–148 in T. Breithaupt and M. Thiel, editors. Chemical communication in crustaceans. Springer, New York. Scholtz, G. 1995. Head segmentation in Crustacea: an immunohistochemical study. Zoology 98:104–114. Scholtz, G. 1997. Cleavage, germ band formation and head segmentation: the ground pattern of the Euarthropoda. Pages 317–331 in R.A. Fortey and R.H. Thomas, editors. Arthropod relationships. Chapman & Hall, London. Scholtz, G., and S. Richter. 1995. Phylogenetic systematics of reptantian Decapoda (Malacostraca: Crustacea). Zoological Journal of the Linnean Society 113:289–328. Seitz, R., K. Vilpoux, U. Hopp, S. Harzsch, and G. Maier. 2005. Ontogeny of the Marmorkrebs (marbled crayfish): a parthenogenetic crayfish with unknown origin and phylogenetic position. Journal of Experimental Zoology 303A:393–405. Shabani, S., M. Kamio, and C.D. Derby. 2008. Spiny lobsters detect conspecific blood-borne alarm pheromones exclusively through olfactory sensilla. Journal of Experimental Biology 211:2600–2608. Sigvardt, K.A. 1977. Sensory-motor interactions in antennal reflexes of the American lobster. Journal of Comparative Physiology A 118:195–214. Sintoni, S., K. Fabritius-Vilpoux, S. Harzsch. 2007. The Engrailed-expressing secondary head spots in the embryonic crayfish brain: examples for a group of homologous neurons in Crustacea and Hexapoda? Development Genes and Evolution. 217:791–799 Spears, T., and L.G. Abele. 1997. Crustacean phylogeny inferred from 16s rDNA. Pages 169–187 in R.A. Fortey and R.H. Thomas, editors. Arthropod relationships. Chapman & Hall, London. Spindler, K.D., P. Jaros, and W. Weidemann. 2000. Arthropoda—Crustacea. Pages 243–269 in K.G. Adiyodi and R.G. Adiyodi, editors. Reproductive biology of invertebrates, Vol. 10, Part A, Progress in Developmental Endocrinology, A. Dohrn, editor. John Wiley & Sons, Chichester. Stensmyr, M.C., S. Erland, R. Wallen, P. Greenaway, and B.S. Hansson. 2005. Insect-like olfactory adaptations in the terrestrial giant robber crab. Current Biology 15:116–121. Stowe, S. 1980. Rapid synthesis of photoreceptor membrane and assembly of new microvilli in a crab at dusk. Cell and Tissue Research 211:419–440. Strausfeld, N.J. 2005. The evolution of crustacean and insect optic lobes and the origins of chiasmata. Arthropod Structure and Development 34:235–256. Strausfeld, N.J. 2012. Arthropod brains: evolution, functional elegance, and historical significance. Belknap Press, Harvard University Press, Cambridge, MA. Sullivan, J.M., and B.S. Beltz. 2001. Neural pathways connecting the deutocerebrum and lateral protocerebrum in the brains of decapod crustaceans. Journal of Comparative Neurology 441:9–22. Sullivan, J.M., and B.S. Beltz. 2005. Integration and segregation of inputs to higher-order neuropils of the crayfish brain. Journal of Comparative Neurology 481:118–126. Tautz, J. 1987. Water vibration elicits active antennal movements in the crayfish Orconectes limosus. Animal Behaviour 35:748–754. Tautz, J., W.M. Masters, B. Aicher, and H. Markl. 1981. A new type of water vibration receptor on the crayfish antenna. I. Sensory physiology. Journal of Comparative Physiology A 144:533–541. Taylor, R.C. 1968. Water-vibration reception: a neurophysiological study in unrestrained crayfish. Comparative Biochemistry and Physiology 27:795–805. Taylor, R.C. 1975a. Physical and physiological properties of the crayfish antennal flagellum. Journal of Neurobiology 6:501–519. Taylor, R.C. 1975b. Integration in the crayfish antennal neuropile: topographic representation and multiple channel coding of mechanoreceptive submodalities. Journal of Neurobiology 6:475–499. Tazaki, K. 1977. Nervous responses from mechanosensory hairs on the antennal flagellum in the lobster Homarus gammarus (L). Marine Behaviour and Physiology 5:1–18. Thiel, M. 2011. Chemical communication in peracarid crustaceans. Pages 199–218 in T. Breithaupt and M. Thiel, editors. Chemical communication in crustaceans. Springer, New York.

47

48

David C. Sandeman, Matthes Kenning, and Steffen Harzsch Tierney, A.J., C.S. Thompson, and D.W. Dunham. 1986. Fine structure of aesthetasc chemoreceptors in the crayfish Orconectes propinquus. Canadian Journal of Zoology 64:392–399. Tonini, G., G.M. Edelman, and O. Sporns. 1998. Complexity and coherency: integrating information in the brain. Trends in Cognitive Sciences 2:474–484. Turvey, P., and J.R. Merrick. 1997. Diet and feeding in the freshwater crayfish, Euastacus spinifer (Decapoda: Parasticidae) from the Sydney region, Australia. Proceedings of the Linnaean Society of New South Wales 118:175–185. Vilpoux, K., S. Bisch-Knaden, and S. Harzsch. 2008. Mapping of Engrailed-expressing cells in the embryonic central nervous system of the marble crayfish (Marmorkrebs). Invertebrate Neuroscience 8:177–197. Vilpoux, K., R. Sandeman, and S. Harzsch. 2006. Early embryonic development of the central nervous system in the Australian crayfish and the Marbled crayfish (Marmorkrebs). Development Genes and Evolution 216:209–223. Wachowiak, M., C.E. Diebel, and B.W. Ache. 1996. Functional organization of olfactory processing in the accessory lobe of the spiny lobster. Journal of Comparative Physiology A 178:211–226. Wirkner, C.S., and S. Richter. 2010. Evolutionary morphology of the circulatory system in Peracarida (Malacostraca; Crustacea). Cladistics 26:143–167. Zeil, J., and J.M. Hemmi. 2006. The visual ecology of fiddler crabs. Journal of Comparative Physiology A 192:1–25. Zeil, J., R. Sandeman, and D. Sandeman. 1985. Tactile localisation: the function of active antennal movements in the crayfish Cherax destructor. Journal of Comparative Physiology A 15:607–617. Zeil, J., G. Nalbach, and H.-O. Nalbach. 1986. Eyes, eyestalks and the visual world of semi-terrestrial crabs. Journal of Comparative Physiology A 159:801–811. Zeil, J., G. Nalbach, and H.-O. Nalbach. 1989. Spatial vision in a flat world: optical and neural adaptations in arthropods. Pages 123–137 in R.N. Singh and N.J. Strausfeld, editors. Neurobiology of sensory systems. Plenum, New York. Zimmer-Faust, R.K., J.E. Tyre, and J.F. Case. 1985. Chemical attraction causing aggregation in the spiny lobster, Panulirus interruptus (Randall), and its probably ecological significance. Biological Bulletin 169:106–118. Zimmer-Faust, R.K., and E. Spanier. 1987. Gregariousness and sociality in spiny lobsters: implications for den habitations. Journal of Experimental Marine Biology and Ecology 105:57–71.

3 SENSORY SYSTEMS OF CRUSTACEANS

DeForest Mellon, Jr.

Abstract Crustaceans, like other metazoans, are endowed with multiple sensory systems that ensure their survival in the presence of predators, and as competitors for resources and mating partners. In most crustacean genera, the chemical senses may constitute the most critical sensory system in terms of their importance in resource detection, reproductive success, and predator awareness. Furthermore, many large crustaceans can live for decades, and a continuously enlarging central olfactory system, via neurogenesis, may permit the consolidation of memories of novel odorants. Tactile and hydrodynamic mechanoreceptors often are coactivated with chemical sensors and the possibility for central integration of these two crucial modalities should be expected. Hydrodynamic sensors, furthermore, are critical for triggering behaviors promoting escape from attacking predators, especially under conditions of low ambient light or poor visibility in the surrounding water column. Crustaceans also possess cuticular and noncuticular proprioceptors for postural maintenance, coordinated locomotion, and visual stabilization. Vision is most highly evolved, and a critical sense in social interactions, among stomatopod crustaceans and certain terrestrial or semiterrestrial crabs; it is also important in triggering defensive behaviors in diurnal decapods. Some decapods exhibit homing and other navigational capabilities, apparently by sensing the Earth’s magnetic field, although the sensory apparatus responsible for this capability has not been identified. Freshwater crayfish have demonstrated a sensitivity to strong electrical fields, but it remains unknown whether this sensory capacity is normally important for prey detection.

O n e c o u l d a l m o s t define a sensory system as a mechanism for throwing away information. —David Young, Nerve Cells and Animal Behaviour

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DeForest Mellon, Jr.

INTRODUCTION Ostensibly, sensory systems act as restricted cellular portals through which selectively critical environmental energy modalities and their dynamic changes initiate or modify behavioral routines. This statement should not imply that sensory systems slavishly and indiscriminately transmit all contingent information that could be conveyed to the central nervous system (CNS) via any particular channel. More often as not, as suggested by Young’s statement above (Young 1989), sensory systems can present significant barriers to specific spectral features of any class of stimulus energy, filtering out the irrelevant, while amplifying and sharpening those bands or qualities crucial to an animal’s lifestyle. Selective filtering starts with the structural properties of the sense organs themselves, and intensifies dynamically through interactions between higher-order sensory neurons that engage in further pruning of stimulus aspects not necessary for an animal’s acute or reproductive survival. In this chapter, I  present an overall review of these adaptive aspects of crustacean sensory systems in selected taxa, against the background of their environmental and behavioral contexts. Crustacean sensory systems, as with most other major metazoan groups, exhibit wide variations on common themes. I will examine all of the major sensory system modalities found across crustaceans, emphasizing common structural and functional designs as well as unique deviations from the usual plan where they occur. By design, none of the systems discussed will be examined in depth, as that responsibility properly falls to the authors of the succeeding chapters on individual specific sensory systems. I hope, however, that the present chapter will serve as a sufficient, general introduction to those chapters. Evidence for more unusual sensory modalities, such as geomagnetic and electrical senses, will be briefly examined, as will the possible ability to detect changes in hydrostatic pressure, and the acoustic far-field.

THE CHEMICAL SENSES I f i n d o u b t, Meriodoc, always follow your nose.—Gandalf — The Fellowship of the Ring

To the extent that humans live in a sensory world dominated by sight and sound, the primary environmental contexts on which the majority of crustaceans depend, both in the aquatic and the terrestrial realms, are chemicals:  food sources, potential mating partners, predators, and even conspecifics with whom an individual has had a recent agonistic encounter, all are primarily recognized through chemicals (Derby and Atema 1982, Devine and Atema 1982, Johnson and Atema 1983, Weissburg and Zimmer-Faust 1994, Moore and Grills 1999, Breithaupt and Eger 2002, Grasso and Basil 2002, Johnson and Atema 2005, Berry and Breithaupt 2010, ­chapter  10 in this volume). To this end, crustaceans are equipped with extensive arrays of cuticular chemoreceptor sensilla. It is reasonable to recognize two major chemosensory systems in crustaceans:  the distributed chemical sense and the olfactory sense (Schmidt and Mellon 2011). Inputs from these two major divisions target entirely different regions of the CNS and in most cases appear to also be separated by threshold ranges, with the olfactory system being more sensitive. In many if not most instances, the distributed chemical sense is mediated through bimodal cuticular sensilla that harbor dendrites of both chemoand mechanosensory neurons. As a class, the distributed chemical sensilla differ in this way from olfactory sensilla, or aesthetascs, which harbor dendrites of exclusively chemosensory neurons.

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Fig. 3.1. Some crustacean bimodal sensilla. (A) Scanning electron microscope (SEM) image of a lateral antennular flagellum from Procambarus clarkii indicating the numerous beaked sensilla (white asterisks) and aesthetasc sensilla (arrowheads). (B) SEM image of a beaked sensillum, with the tip (inset) at higher magnification. (C) Transmission electron microscope (TEM) image of a transverse section through the cuticular canal near the base of a beaked sensillum. Asterisks indicate putative distal dendrites of mechanoreceptor neurons (see text), adjacent to transverse sections through five distal dendrites of presumed chemoreceptor neurons. A scolopale (SC) surrounds this section of the distal dendrites, and all are enclosed in a sheath composed of enveloping cells (EC). (D) TEM image of distal dendritic segments through the cuticular canal at the base of a hedgehog bimodal sensillum from the crayfish Austropotamobius torrentium. At this level, the distal dendrites are surrounded by a cuticular sheath (CS). There are sections through six putative chemoreceptor distal dendritic segments and two putative mechanoreceptor distal dendritic segments (asterisks) (from Altner et al. 1983, with permission from Springer). (E) SEM image of a hedgehog sensillum from the chela of a first pereopod of Homarus americanus (from Derby 1982, with permission from BioOne). (F) SEM image of a hooded sensillum from the lateral antennular flagellum of Panulirus argus. The distal end of the central shaft of the sensillum (single white arrow) is serrate and is partially surrounded by a hood composed of fused setules (double headed arrow). (G) TEM image through the outer dendritic segments of a hooded sensillum from P. argus in the region surrounded by a scolopale (as indicated on the figure). Arrows indicate sections through three dendrites from putative mechanoreceptor neurons. Approximately 10 additional (presumably chemoreceptive) dendrites are visible in the section. microtubules (mt). (F and G from Cate and Derby 2002, with permission from John Wiley and Sons). Scale bars: A, 100 μm; B, 1 and 5 μm; C, 1 μm; D, 1 μm; F, 10 μm; G, 0.2 μm.

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DeForest Mellon, Jr. Distributed Chemical Sense Distributed chemical sensilla in crustaceans are found on the flagella of the first antennae (antennules), mouthpart appendages, including the maxillipeds, maxillae, and mandibles, and the propodus and dactyl of the pereopods. Bimodal sensillar structures vary widely not only among different groups but also within individual species, ranging from comparatively simple setae, such as the smooth or beaked sensilla of the antennal and antennular flagella in the crayfish Procambarus clarkii and the asymmetric sensilla of the lateral antennular flagellum in the spiny lobster Panulirus argus, to more elaborate structures, such as the hooded sensilla found in Panulirus and the hedgehog hairs of the pereopod dactyls in crayfish and the lobster Homarus americanus (see Fig. 1 in Derby 1989). The asymmetric sensilla in Panulirus are noteworthy in that they are uniquely responsible for triggering the antennular grooming reflex when stimulated with the amino acid L-glutamate (Schmidt and Derby 2005). A sensillar type on the propodus and dactyls of decapod walking legs, called funnel-canal organs, are instrumental in probing for food on and within the substratum, a behavior that is vigorously employed in the presence of appropriate chemical stimuli within the water column. Other bimodal sensilla on the pereopods include the hedgehog sensilla of crayfish and lobster chelae. The electrophysiological records in Fig. 3.2 illustrate characteristic responses to chemical and mechanical stimulation of different types of chemo- and bimodal sensilla of decapods. As typified by those described for the crayfish Austropotamobius torrentium (Fig. 3.1D), the hooded sensilla of the spiny lobster antennules (Fig. 3.1F, G), the sensory neuron dendrites that span the cuticle and supply these sensilla are of two major types that, as discussed below, appear to subserve, respectively, chemoreception and mechanoreception functions of the sensillum. The dendrites are enclosed in a dendritic sheath, both within the hypodermis and the cuticular canal. Moreover, the sheath is enveloped in supporting cells that insulate it from the wall of the canal. Using the above fine structural criteria, beaked sensilla on the medial and lateral flagella of the crayfish P. clarkii may also be presumed to have a bimodal function. Transmission electron microscopy of transverse sections of these sensilla proximal to their base within the cuticle reveals large-diameter outer dendritic segments with a high packing density of microtubules and more numerous, smaller diameter dendrites with low microtubule density. A sleeve of osmophilic tissue referred to as the scolopale tightly surrounds the transitional region between the inner and outer dendritic segments (Fig. 3.1C). This is a characteristic of cuticular mechanoreceptor sensilla in other arthropod groups as well (e.g., Gray 1960, Altner et al. 1983, Schmidt and Gnatzy 1984, Derby 1989, Cate and Derby 2002, Schmidt and Derby 2005). Behavioral observations with crayfish suggest that contact chemoreceptors on the antennular flagella are used to identify possible food sources, since these animals will often depress their antennules to touch a potential food item on the substratum. Bimodal, as well as strictly mechanosensitive, setae have been described on the antennules of the copepod Pleuromamma xiphias, based on the fine structure observed in transmission electron micrographs (Weatherby et al. 1994). Copepod antennules are large and elaborate and, from behavioral observations, they are finely tuned to detect chemical as well as hydrodynamic stimuli in the environment (Katona 1973, Gill and Crisp 1985). From physiological studies, calanoid antennular setae have been found to have an order of magnitude higher sensitivity to mechanical stimuli than comparable receptors on decapods (Yen et al. 1992, Lenz and Yen 1993). On the pereopods of both Homarus and Panulirus, as well as the crayfish, bimodality of contact sensilla is the rule; here extracellular electrophysiological data (see Fig. 3.2) indicate that spike amplitudes, and hence, axon diameters, are larger for the mechanosensory neurons than for the chemosensory neurons (Derby 1982, Altner et al. 1983). This assumption is supported by ultrastructural analysis where comparisons of the outer dendritic segments of mechano- and

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Fig. 3.2. Physiological responses of sensory neurons. (A)  Spiking response from a chemoreceptor neuron in a hedgehog sensillum in the crayfish Austropotamobius torrentium following exposure to 10 -4 mol/l serine. Solid line beneath the electrical record indicates period of serine application. Time calibration applies to both A and B (from Altner et al. 1983, with permission from Springer). (B) Spiking responses from two mechanoreceptor neurons in a crayfish hedgehog sensillum different from that in A, to mechanical displacements; both mechanoreceptive units of the sensillum respond to three stimulations, the durations of which are shown by the solid lines beneath the records. (C) Spiking responses of a chemoreceptor neuron in a hooded sensillum fromn Panulirus argus to artificial sea water (ASW) and to shrimp extract. st, stimulus. In (D), a mechanoreceptor neuron in a different hooded sensillum than in C responded with a phasic spike discharge to mechanical depression, indicated on the lower trace (from Cate and Derby 2002, with permission from John Wiley and Sons). (E) Whole cell patch clamp recording of voltage changes in an olfactory receptor neuron from P. argus to application of tetramarin (a food stimulus) to the aesthetasc housing its dendrites. Reduction in spike amplitude during the record is the result of the intensity of stimulation (from Schmiedel-Jakob et al. 1989, with permission from The American Physiological Society). (F) Patch clamp recording of currents from a single histamine-gated ion channel in an olfactory receptor neuron of P. argus. Open (O) and closed (C) positions of the channel are indicated (from McClintock and Ache 1989, with permission from the National Academy of Sciences).

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DeForest Mellon, Jr. chemosensory neurons within the same sensillum exhibit, respectively, larger overall diameters, greater numbers, and a higher packing density of microtubules, and where the presence of mechanosensory dendrites can be specifically suspected by the existence of a scolopale (Cate and Derby 2002, Schmidt and Derby 2005). The presence of bimodal sensilla on the second antennae of decapods is somewhat problematic. Ball and Cowan (1977) suggest that one type of seta on the antennae of the sergestid shrimp Acetes may have a chemoreceptive, as well as a mechanoreceptive function. The lumen of the seta contains up to 10 outer dendritic segments, and there is a definite pore at its tip. There is also a scolopale at the transition region, indicating, as suggested above, the presence of mechanoreceptor function as well. Reports of chemo- and mechanoreceptor sensilla on the antennae of nondecapod malacostracans have appeared, though infrequently. Guse (1978) has detailed antennal sensilla in the mysid Neomysis integer. The fine structure of the inner and outer dendritic segments again suggests the presence of both chemoreceptor and mechanoreceptor neurons associated with these setae. Alexander (1977) described antennal sensilla in the isopod Ligia oceanica using both ultrastructural and electrophysiological techniques to determine their functional properties. The setae are short, hooded structures containing several outer dendritic segments with microtubules that terminate near the tip. Each seta has only one sensory neuron associated with it. Recordings from the antennal nerve reveal spikes in response to contact with the substratum or other solid objects; in addition, however, a withdrawal response of the antenna occurs in intact animals following application of L-glutamate to the tips of the sensilla. It is unclear whether the response to mechanical stimulation actually originated from the sensilla cluster or from other undetected mechanoreceptors. No scolopale was mentioned in the ultrastructural description of these sensilla. In P. clarkii, beaked sensilla similar in form to those found on the antennules are prevalent on the antennae. If they are similar in their function to that alleged in the case of the antennular sensilla, they would convey chemical as well as mechanical sensitivity to the antennae. The central targets of bimodal sensilla depend critically on the peripheral location of the latter. Those on the pereopod chelae presumably target circuits within the appropriate thoracic ganglia that reflexively control grasping and transfer of materials to the mouth. In crayfish, and presumably in other macruran decapods, objects grasped by the chelae of the pereopods are smoothly transferred to the maxillae even in the absence of a brain (Huxley 1896, Bethe 1897). Chemo- and mechanoreceptor afferents on the maxillae, maxillipeds, and mandibles enter the subesophageal ganglion, where they drive or influence activity in the motor neurons to those appendages. It is possible (but currently unknown) that collateral branches of mouthpart chemoreceptor afferents ascend to the brain, where their information may be integrated with chemical input from the antennules. Afferent inputs from bimodal sensilla on the antennules of the spiny lobster run within the antennular nerve primarily to the lateral antennular neuropils within the deutocerebrum and possibly, but if so to a lesser extent, to the median antennular neuropil (Schmidt et al. 1992). These regions are invested with the dendrites of motor neurons that control antennular movements and thus are undoubtedly sites for synaptic interactions driving reflex movements of the antennules and/or antennae (e.g., Schmidt et al. 1992, Mellon 1997). Antennal afferents target postsynaptic sites within the striated neuropil of the tritocerebrum, the termination point of the antennal nerve, in what appears to be a topographic fashion (Schmidt and Mellon 2011), presumably thereby being capable of localizing the site of stimulation on the antenna. Additional target neuropil served by nascent afferent terminals is added as the animal grows. From the tritocerebrum, information about mechanical and sensory input may spread to other brain regions that integrate broad spectrum sensory inputs, such as the accessory lobes (Sandeman et al. 1995) and the hemiellipsoid bodies of the lateral protocerebrum (Mellon 2000).



Sensory Systems of Crustaceans

Olfactory Sense The olfactory sense of crustaceans is mediated by cuticular sensilla referred to as aesthetascs. These structures are arrayed along the ventral surface of the lateral antennular flagellum, grouped in varying numbers and spacing density on each flagellar annulus in a species-specific manner. Depending on species, aesthetascs are 100–600 μm in length and at their base range in diameter from 10 to 20 μm (Fig. 3.3A–D). They are not characterized by microscopically visible pores at their tips or elsewhere along their length, although their cuticle is extremely thin at around 1 μm. In crayfish, the distal 40% of the aesthetasc shaft is optically transparent and is coincident with the region that is permeable to dissolved aqueous solutes (Tierney et al. 1986). The ultrastructure of decapod aesthetascs is interesting and can be typified by that of crayfish aesthetascs (Tierney et al. 1986). Each aesthetasc contains the outerdendritic segments of up to several hundred olfactory receptor neurons (ORNs), whose cell bodies are grouped in a series of clusters arrayed within the lateral flagellar axis near the base of each of their parent aesthetascs. After the dendrites emerge from their respective somata, each divides in two. The dendrites remain in this configuration as they course distally, first passing through a fluid-filled cavity known as the receptor lymph cavity before becoming tightly wrapped in multiple layers of supporting, or sheath, cells. This region of the outer dendrites and their sheath course through another fluid-filled cavity, the outer lymph cavity, which is itself surrounded by layers of outer supporting cells (Fig. 3.3E). Finally, as they enter the lumen of the aesthetasc and continue toward its tip, the outer dendritic segments subdivide into thousands of individual microtubules. Our current understanding of the functional properties of aesthetascs is that olfactory receptor proteins are embedded in the microtubular membranes (Hatt and Ache 1994). The receptor proteins from each ORN bind only a single type of olfactory determinant, possibly one of many types of atomic moieties found on an odorant molecule. Each aesthetasc is thus considered to be a repeat unit, housing identical families of ORN receptors. The combinations of olfactory determinants detected by subsets of ORNs on exposure to odorant molecules, and which uniquely define that molecule, are presumably sorted out within the brain. The distribution, arrangement, and number of aesthetascs on the antennules of different crustacean species varies unpredictably across taxa, and in species with indeterminate growth, their numbers show a net increase as the animal grows and the antennules become longer. A second posthatch instar of the crayfish P. clarkii, for example, may have 3–5 aesthetascs on each lateral flagellum, whereas a large adult of 70 mm carapace length can have more than 160 per flagellum (Mellon et al. 1989). Growth and the appearance of new sensilla occur at or near the flagellar base (Sandeman and Sandeman 1996). The conventional wisdom is that increased numbers of identical sensors increases the signal-to-noise ratio within the CNS processing centers. In terrestrial anomurans, such as Coenobita and Birgus, the aesthetascs have undergone morphological changes that cause them to resemble insect olfactory sensilla more than those of closely related marine hermit crabs. The individual aesthetascs are short and stubby, and electron micrographic examination of the aesthetascs in Coenobita and Birgus (Ghiradella et al. 1968, Stensmyr et al. 2005, respectively) reveals them to have an asymmetrical structure; the surface cuticle of that region of the aesthetasc facing the antennule is heavy and thick, while the cuticle facing the environment is thin and crenulated, possibly as a mechanism to increase surface area. These findings suggest that convergent evolutionary adaptations have occurred in terrestrial crustaceans from selective pressures that uniquely involve differences between the aquatic and terrestrial environments. Due to the recently confirmed presence of distributed (bimodal) sensilla on the antennular flagella of spiny lobsters (e.g., Cate and Derby 2002), much of the early electrophysiological data on aesthetasc sensitivity are suspect, since, with extracellular recordings from chemoreceptor

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Fig. 3.3. Aesthetasc sensilla. (A) Light micrograph of a portion of a living lateral antennular flagellum of Procambarus clarkii. Arrows indicate sparse distribution of pairs of ventrally located aesthetascs on annuli along the outer one-half of the flagellum. (B) Nomarski optics light micrograph of a portion of a living lateral antennular flagellum from P. clarkii. White asterisk indicates a cluster of cell bodies of olfactory receptor neurons associated with the aesthetasc indicated by the arrow, visible through the semitransparent cuticle. The white line is the approximate position of the section through an aesthetasc’s outer dendritic segments shown in E. (C) Light micrograph showing array of aesthetascs (arrows) on an antennular flagellum of the marine hermit crab Pagurus bernhardus. (D) shows an SEM image of clusters (arrow) of the individual aesthetasc sensilla at higher magnification. The dense spacing of the sensilla and their filiform external morphology are in stark contrast to the situation in the crayfish (modified from Hansson et al. 2011, with permission from Springer). (E) TEM image of a section through the outer dendritic segments of approximately 175 olfactory receptor neurons below the base of an aesthetasc from P. clarkii. The outer dendrites are tightly wrapped by enveloping inner sheath cells (ISC). OLC is the outer lymph cavity, and OSC is the outer sheath cell wrapping the entire structure (from Mellon et al. 1989, with permission from John Wiley & Sons). Scale bars: A, 250 μm; B, 100 μm; E, 1 μm.



Sensory Systems of Crustaceans

axons at the flagellar base, it is very difficult, if not impossible, to determine the sensillar origin of chemically evoked spikes. Since the late 1980s however, patch clamp technology has been used to examine the response characteristics of identified ORNs in P. argus antennular preparations (Anderson and Ache 1985, McClintock and Ache 1989, Schmiedel-Jakob et al. 1989), from cultured ORN cell bodies (Fadool et al. 1993), and even from vesiculated membrane patches from ORN outer dendrites (Fadool and Ache 1992, Hatt and Ache 1994). With only a few exceptions, most of what we currently understand about the response properties of crustacean ORNs and their signal transduction pathways have been derived from studies with this animal. Lobster ORNs are sensitive to a variety of amino acids, complex odorants such as tissue extracts, and amines such as histamine. Olfactory determinants bind to receptors in lobster ORNs and initiate signal transduction cascades that result in opening of ion channels, leading either to excitation or inhibition. Excitation is mediated by inositol 1,4,5-trisphosphate-activated channels (Fadool and Ache 1992), whereas inhibition depends on cyclic nucleotide-gated channels (Hatt and Ache 1994, Bobkov et al. 2010). Spike responses to acceptable odorants occur as tonic spike trains exhibiting concentration-dependent frequency modulation, or, in about 30% of ORNs, intrinsically generated spike bursts whose probability of being triggered is also odorant concentration-dependent (Bobkov and Ache 2007). In all crustaceans, ORN axons run directly to targets in the ipsilateral olfactory lobes (OL) of the deutocerebrum (Sandeman and Denburg 1976, Mellon et al. 1989, Schmidt et al. 1992). The neuropil of the OL is highly structured, being compartmentalized into columnar glomeruli, in each of which information arriving from one class of olfactory determinant-specified ORNs is processed. In crayfish, and probably in other crustaceans, each aesthetasc accommodates the dendrites of all ORN classes, since labeling of a single aesthetasc with tritiated leucine, which presumably is taken up by all ORN dendrites, appears in all OL glomeruli (Mellon and Munger 1990). Within the glomeruli, ORN axon terminals make synaptic connections with local interneurons and with projection neurons, the latter transmitting locally processed information to neural centers in the lateral protocerebrum, in crayfish including especially the terminal medulla, and possibly the hemiellipsoid body (Wachowiak and Ache 1994, Mellon and Alones 1995, Schmidt and Ache 1996, Sullivan and Beltz 2001, 2004, 2005). Electrical recordings and morphological studies employing dye filling from local OL interneurons in crayfish and spiny lobsters indicate that they are broad-spectrum, receiving input via their dendrites from many if not all of the OL glomeruli (Mellon and Alones 1995, Schmidt and Ache 1996). At least two classes of OL local interneurons in crayfish receive hydrodynamic input as well, via dendritic branches within the lateral antennular neuropil (Mellon 2005). In most, but by no means all decapods, the OL are intimately connected with another large, glomerular deutocerebral center for integration of multimodal sensory input, the accessory lobes (AL) (Sandeman et al. 1992, Sandeman et al. 1995, Wachowiak et al. 1996). Where it has been critically examined, the organizational plan of glomeruli within the AL is characterized by a segregation into medullary and cortical zones, with higher-level processing of tactile and visual inputs occurring in the glomeruli of the medulla, and input from the OL being processed in the cortex (e.g., Sullivan and Beltz 2005). Output from the AL is also transmitted via projection neurons to the hemiellipsoid bodies within the lateral protocerebrum (Sullivan and Beltz 2001). Attempts to formulate a generalized statement regarding specific details of the higher olfactory pathway in malacostracan crustaceans are complicated both by the considerable variability across taxa in the structural organization of the hemiellipsoid bodies and the reduction or absence of accessory lobes in some groups (e.g., Sullivan and Beltz 2004). Of recent and compelling interest is that cellular groups within the olfactory pathway of decapods exhibit lifelong neurogenesis, especially among the OL projection neurons and the local interneurons in crayfish and lobsters, where it has been most intensively studied (Schmidt 2007,

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DeForest Mellon, Jr. Sullivan et al. 2007, ­chapter 7 in this volume). It has been argued (Schmidt and Mellon 2011) that continued generation of nascent projection neurons enable these relatively long-lived decapods to acquire novel neural pathways via combinations of glomerular inputs that correspond to new odorants the animals will experience within their environment throughout their lives.

THE ELECTROMAGNETIC SENSES A l l t h a t f o l l o w their noses are led by their eyes but blind men. —William Shakespeare, King Lear, II, iv

Like all other arthropods, crustaceans are endowed with several kinds of light-sensing peripheral organs and photoreceptive neurons in the CNS. Photosensitive neurons are found within the brain (Sandeman et al. 1990, Martin et al. 1995, Sullivan et al. 2009) and in the last abdominal ganglion of adult crayfish (Larimer 1966). Most crustaceans also have a median, or nauplius eye, at least in their early larval stage, and the majority of adult malacostracans have lateral compound eyes, highly structured image-forming organs, many of which are comparable to or exceeding those found in insects in their acuity, spectral sensitivity, and resolution capabilities. Extraretinal Photoreceptors Extraretinal brain photoreceptor neurons were first discovered in the parastacid crayfish Cherax destructor by Sandeman et al. (1990) (see Fig. 3.4), although the existence of photosensitivity in the crayfish nervous system was suspected earlier following the discovery that locomotory activity could be photically entrained following removal of the compound eyes as well as the caudal photoreceptor in the sixth abdominal ganglion (Page and Larimer 1976). Studies on the brain photoreceptors in C. destructor (Sandeman et al. 1990) and P. clarkii (Sullivan et al. 2009) have shown that brain photoreceptor neurons exist as bilateral clusters close to the midline and at the dorsal-anterior surface of the protocerebrum. A portion of the apposed plasma membranes of the component cells within a cluster is made up of clustered microvilli, which collectively form a rudimentary rhabdom (Fig. 3.4B). The cells react with an antibody to the blue light-sensing photopigment cryptochrome; furthermore, as shown earlier by Page and Larimer (1976), locomotory activity rhythms can be photically entrained as well as phase shifted in animals lacking both functional compound eyes and caudal photoreceptors, implicating the brain photoreceptors as the zeitgeber. The axons of brain photoreceptor neurons terminate in paired protocerebral neuropils among neural processes expressing pigment-dispersing hormone, a substance thought to be involved in circadian signaling in crustaceans (Sullivan et al. 2009). Brain photoreceptor neurons have also now been discovered in the giant isopod Glyptonotus antarcticus (Martin et al. 1995). The caudal photoreceptor of the sixth abdominal ganglion in the crayfish was discovered by Prosser (1934) and studied more methodically by Kennedy (1958, 1963). It is found in both epigean and cavernicolous crayfish species (Larimer et  al. 1966). The caudal photoreceptor’s sensitivity to illumination is confined to (presumably the cell bodies in) the sixth abdominal ganglion and comprises two neurons whose axons run in a rostral direction within the ventral nerve cord. The activity of the photically excited axons is characterized by a steady train of impulses whose frequency and latency depend critically on stimulus intensity (Kennedy 1958). Early behavioral work by Welsh (1934) showed that its activation was responsible for backward walking as well as flexion of the abdominal segments. These experiments were revisited by



Sensory Systems of Crustaceans

Edwards (1984), who used electrophysiological analysis of motor neuron output to examine the links with caudal photoreceptor activation. Kennedy (1963) showed that the neurons of the caudal photoreceptor are also mechanoreceptor interneurons that have a low threshold for and are excited by low intensity ipsilateral mechanoreceptor afferent inputs and are inhibited by higher intensities of stimulation. Contralateral afferent inputs cause only inhibition of the photoreceptor neurons. It was suggested that a functional significance of the caudal photoreceptor inhibitory inputs might be detection of weak or protracted mechanical disturbances in the water near the tail fan (Kennedy 1963, see also review by Wilkens 1988). A search of the literature reveals no ultrastructural studies of the crayfish caudal photoreceptor. Hama (1961) described structures associated with the giant axons in the crayfish abdominal nerve cord as having a structure resembling “known types of photoreceptors in animals and plants,” but their anatomical distribution within the ventral nerve cord casts doubt on any association with the physiologically identified caudal photoreceptor. Nauplius (Simple) Eyes Almost all subclasses of Crustacea possess at least nauplius (i.e., median) eyes in the larval state, although even these have been secondarily lost in some malacostracan orders (Amphipoda, Isopoda). A review of nauplius eye distribution can be found in Waterman (1961), and a comprehensive comparative morphology of nauplius eyes is found in Vaissiere (1961). The nauplius eye consists of 3–7 tightly apposed ocelli, each of which is an epithelial cup containing as many as ten individual retinula cells. All copepods possess a median nauplius eye, and it can serve adequately as a typical example of the general structure. Fahrenbach (1964) described the fine structure of the nauplius eye in the copepod Macrocyclops albidus, diagramed in Fig. 3.4D. The proximal region of the eye cup is lined with a hemispherical tapetum formed by two specialized cells and encloses nine irregularly shaped retinula cells. Each retinula cell has a portion of its plasma membrane thrown into tightly grouped microvilli—the rhabdomere—the membranes of which contain the photoreceptor pigment. The rhabdomeres of each retinula cell are oriented in such a way that the microvilli are orthogonal to the direction of light rays. Moreover, as in the compound eyes of crustaceans and other mandibulate arthropods, all of the rhabdomeres are contiguous and form a rhabdom. The most conspicuous crustacean simple eyes belong to the ostracod Gigantocypris, a deep-sea, spherically shaped species that can reach 30 mm in diameter. Gigantocypris is notable for its huge paired ocelli, backed by a highly efficient parabolic mirror-tapetum that focuses light on the rhabdom. As these animals are predators of other deep-sea plankton and nekton at depths far below the penetration of any incident sunlight, it is thought that the eyes provide a means for detecting bioluminescent prey. Compound Eyes With the exception of the Phyllocarida, all subclasses of the Malacostraca have compound eyes, some of which are structurally and functionally highly complex. Among the extant Hoplocarida—the stomatopods---- the compound eyes are capable of excellent, multiwavelength color vision and also possess the ability to detect both plane- and circularly polarized light (see ­chapters 8 and 9 in this volume). These and the eyes of various decapods have been most intensively studied. Both apposition and superposition compound eyes are found in decapod malacostracans. In crustacean apposition compound eyes, as in those of insects, the cornea acts as a lens to focus light rays through a long lens cylinder, or crystalline cone, the refractive index of which decreases in an essentially parabolic gradient outward from the central optical axis (Exner 1891). Light entering an ommatidium is thereby focused directly on the rhabdom of

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Fig. 3.4. Crustacean photoreceptors. (A) Light micrograph of a cluster of brain photoreceptor neurons from the crayfish Cherax destructor. The neurons contain visible dark pigment granules and have axons that terminate in the protocerebrum. Scale bar: 20 μm. (B) TEM image of three photoreceptor cells (numbered) from the brain photoreceptor in Cherax. Round dark pigment granules are prominent; a component of each cell joins with others to form a crude rhabdom (Rh), shown at greater magnitude in the inset. Scale bars 5 μm and 1 μm (inset). (C) Intracellular records (upper traces) from a brain photoreceptor neuron in response to light flashes (lower traces) of 1-sec duration during (a–b) dark adaptation. c shows the response after complete dark adaptation. Very low levels of light generate quantum bumps on the response (d). Voltage scale: a, b, 10 mV; d, 5 mV (from Sandeman et al. 1990, with permission from John Wiley & Sons). (D) Drawing of an ocellus from the nauplius eye of the copepod Macrocyclops albidus showing three of the retinula cells (R) and their rhabdomeres (rh) that comprise the rudimentary rhabdom surrounding the dendrite of an eccentric cell (ec). A bowl-shaped tapetum (T) is at the back of the ocellus. Retinula cell axons (ra) course to the brain (modified from Fahrenbach 1964, with permission from Springer). (E) Intracellular records from a cell in the compound eye of the isopod Ligia to increasing levels of illumination (duration of light stimulation indicated by horizontal bars). At high light levels, a distinct initial transient phase of the response can be seen, followed by a more sustained plateau of depolarization (from Shaw and Stowe 1982, with permission from Academic Press).



Sensory Systems of Crustaceans

the underlying, apposed retinula cells, the distal margins of which generally abut the crystalline cone. In superposition compound eyes, the crystalline cones and retinula cell borders are physically separated from each other by a so-called clear zone. Euphausid and mysid shrimp have refracting superposition eyes. In these, as in all superposition eyes, several ommatidia cooperate to form a focused image on individual rhabdoms. While in the apposition eyes of crustaceans the focal length of the crystalline cone and its length are equal, and light rays are thus focused at its base as an inverted image (Fig. 3.5A1), in refracting (and other) superposition eyes, the length of the crystalline cone is twice the focal length of the lens cylinder. The result is a recollimated upright image at the rhabdom from rays along the optical axis and from adjacent, off-angle ommatidia by virtue of the graduated index of refraction in the crystalline cones (Fig. 3.5A2,B). Two other kinds of optical arrangement are found as well in the superposition compound eyes of decapods: reflecting superposition eyes (Land 1976, 1980, Vogt 1977) and parabolic superposition eyes (Nilsson 1988). In the reflecting superposition eyes, found in freshwater crayfishes, lobsters, and many shrimps, the ommatidial facets are square in transverse section (Fig. 3.5C), and the four sides of the tapering crystalline cone act as mirrors that are at right angles to one another. Light entering an ommatidium at an angle to the optical axis is refracted by the cornea and reflected from one or two walls of the crystalline cone before being focused as an upright image on a rhabdom (Fig. 3.5A3). The most optically complex decapod superposition eyes are those found so far only in crabs and hermit crabs. This is the parabolic superposition eye first described by Nilsson (1988) and employing at least four optical mechanisms within each ommatidium. The cornea and crystalline cone cross sections are usually round; the cornea focuses light incident on the optical axis as an upright image on a light guide that begins at the base of the crystalline cone and ends in an on-axis rhabdom. Off-axis light rays are reflected, and their convergence reversed, by the inwardly bulging parabolic walls of the ommatidium’s crystalline cone, then are recollimated by a cylindrical lens within the crystalline cone and finally focused at an appropriate rhabdom, again as an upright image (Fig. 3.5B). In a few parabolic superposition eyes (e.g., xanthid crabs), there is no internal cylindrical lens; instead, the cross section of the parabolic crystalline cone walls is square, and these eyes employ the same focusing principle as the purely reflecting superposition eyes. The advantage of superposition eyes when compared with those of the apposition types is primarily in their greater light-capturing power due to ommatidial cooperativity, and it is usual to find crustaceans having superposition eyes at their most active in crepuscular or nocturnal environments. Resolving power may also be better than that found in apposition eyes, through the intervention of complex optical mechanisms mentioned above. Compound eyes in general, however, all suffer from image degradation due to diffraction, an inherent problem with small-aperture lenses, since diffraction is inversely proportional to aperture diameter. In this sense, therefore, superposition decapod eyes are on average at no greater advantage than apposition eyes. In all compound eyes, the elongated photoreceptor, or retinula, cells within each ommatidium are arranged concentrically around a central axis formed by the abutting or interdigitating rhabdomeres of each. This central composite structure, referred to as the rhabdom, is thus the photosensitive region for each ommatidium. Retinula cell axons course centrally to the underlying lamina where they make synaptic connections with first order visual interneurons. Many, perhaps most decapods, as well as stomatopods, can detect plane-polarized light. Analysis occurs by virtue of the orthogonal arrangement of the microvilli arrays within the rhabdomeres of each of two groups of retinula cells. Figure 3.5D and E shows this arrangement in a generalized crayfish and in the stomatopod Gonodactylus. In most crustacean compound eyes, the rhabdomeres of retinula cells R1, R4, and R5 are arranged orthogonally to those of R2, R3, R6, and R7. The small diameter (0.07–0.08 μm) and hollow structure of the microvilli

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Fig. 3.5. Features of crustacean compound eyes. (A) Three major types of crustacean compound eye optics and their respective focusing mechanisms:  apposition, on the left; refracting superposition, center; and reflecting superposition, on the right (from Land 1980, with permission from Macmillan Publishers). (B) Parabolic superposition optics as discovered by Nilsson (1988) and characterized by parabolic reflecting surfaces within the crystalline cones, light guides for on-axis rays through the clear zone, and with the crystalline cones either containing a cylindrical lens or being square in transverse section (redrawn from Nilsson 1988). (C) SEM image of the surface of the compound eye of the crayfish Procambarus clarkii showing the square crystalline cone facets characteristic of astacid and homarid decapod ommatidia. Each crystalline cone is 55 μm on a side. (D1) Highly simplified diagram of a longitudinal section through a crayfish retinal ommatidium. Two of the eight retinula cells are shown, including the rhabdom, composed of interdigitating microvillar central regions (rhabdomeres) of each of seven of the circularly clustered retinula cells (RC). N, nucleus (redrawn from Eguchi 1999). (D2) Diagram illustrating the mode of rhabdomere interdigitation in neighboring retinula cells (redrawn from Eguchi 1999). (D3) Diagrams of cross sections of an ommatidium at two different levels to illustrate the orthogonal arrangement of the rhabdomeres in neighboring retinula cells. In each ommatidium, retinula cells 1, 4, and 5 give rise to one group of microvillar arrays, while cells 2, 3, 6, and 7 have arrays oriented orthogonally to the first group. (E) TEM image of a longitudinal section through part of a rhabdom of the compound eye of the stomatopod Gonodactylus bredeni. The microvilli of individual retinula cells on the right and the left (indicated by brackets) are arrayed orthogonally to those of the cell whose rhabdomeric microvilli are seen end-on above and below them. D3 and E modified from Eguchi 1999, with permission from Springer.



Sensory Systems of Crustaceans

constrain the positional arrangement of visual pigment molecules within the microvillar wall, which align themselves with their dipole moment in parallel with its long axis. Since the most efficient absorption of electromagnetic radiation is for those rays having their e-vector in parallel with the primary axis of the microvilli, this is thought to provide the basis for the plane polarization analyzer capabilities of most arthropod compound eyes. In some species of the stomatopod genus Odontodactylus, individuals are capable of detecting circularly polarized light, a sensory ability that is believed to be unique in the animal kingdom (Chiou et al. 2008). This capability depends on a modification of one retinula cell, R8, having a distally located rhabdomere, in rows 5 and 6 of the central ommatidial band in stomatopod eyes (Cronin et al. 2000, Chiou et al. 2008). Within these specific ommatidia, R8 acts as a one-quarter-wave plate that translates incident, clockwise or counterclockwise circularly polarized light into plane-polarized light for acquisition by the two sets of orthogonally arranged microvilli in the rhabdomeres beneath it. It is well known that stomatopods use reflected plane-polarized light as social signals. Interestingly, the uropods and telson of some species of Odontodactylus males also reflect circularly polarized light. Thus, this unique sensory ability may have an important role in social interactions of mantis shrimps (Fig. 3.6A, see color version in centerfold) (Chiou et al. 2008). Color vision in crustaceans has been most intensively examined in the stomatopods. These littoral and sublittoral, highly visual cursorial predators can be brightly colored (see Fig. 3.6 color version in centerfold) and apparently use color patterns (as well as the polarization patterns mentioned above) and an excellent capability for color vision in intraspecific and interspecific communication (e.g., Marshall et al. 1999, Cronin et al. 2000, ­chapter 9 in this volume). As many as eight primary classes of retinal photoreceptors based on their respective wavelength absorption spectra, with maxima ranging from 400 to 700 nm, can be found in the compound eyes of some stomatopods, and spectral filtering by pigment plugs within the rhabdoms of different ommatidia increase the diversity of color sensitivities by the receptor cell classes to more than 16 (Cronin et al. 1994). Other than their importance in triggering escape responses to threats from potential predators (e.g., Liden and Herberholz 2008), compound eyes in the crayfish and, presumably, in other decapods are instrumental in stabilizing the visual fields of the retinas through reflex movements of the eyestalks (Schöne 1954, Fay 1973, Mellon and Lorton 1977). Especially, however, crustacean visual systems can be critical in social interactions, as suggested in the previous discussion of stomatopods. Fiddler crabs (Brachyura; Grapsidae) are semiterrestrial, quasi-social decapods living in large colonies within the mudflats and marshes of the intertidal zone, which provide abundant possibilities for sexual and agonistic encounters. While they are foraging, fiddler crabs are vigilant that their burrows do not become occupied by neighboring intruders (see ­chapter 18 in this volume). They calculate the position of the burrow’s entrance and the trajectory and distance from the burrow of potentially invading neighbors, and, if the potential intruder gets too close to a threshold distance from the burrow’s entrance, the owner rushes back to repossess. Remarkably, some species of fiddler crab can infer the position of their burrow entrance by means of path integration, even when it is invisible, and the owner uses vision to determine the threshold distance of a potential intruder to the inferred entrance position at which it must rush back to protect its burrow (Hemmi and Zeil 2003). It has also been shown that fiddler crab (Uca capricornis) males can recognize the carapace color patterns of familiar neighboring female crabs as opposed to strangers, and will more often make mating approaches to the strangers. In Uca moebergi, moreover, females primarily base their recognition of conspecific males on claw color (Detto et al. 2006). Casual observations of the behavior of a number of other semiterrestrial crabs suggest that they are endowed with excellent vision, including binocular depth perception. Grapsus grapsus crabs are denizens of tropical rocky shores, which they traverse with alacrity, even jumping nimbly from one rock to another over distances as great as 0.5 meters (personal observation), a feat

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Fig. 3.6. (A) The stomatopod Odontogonodactylus scyllarus displaying an example of the spectral flamboyance believed to serve as intraspecific communication in these crustaceans (from Debelius 1999, with permission from Ikan). (B) Response-intensity function (curve) for behavioral responses of the crayfish Cherax destructor following stimulation by step voltages applied to the animal’s aquarium. Filled circles are total number of behavioral responses of all animals in the test to 30–50 stimulus presentations at each electric field strength. Open circles represent responses to electric signals generated by swimming animals (indicated by icons) whose signal characteristics are shown below each of the three data points. Scale bar at lower right is 1.5 sec for the tadpole signal, 100 msec for the other two signals (from Patullo and Macmillan 2007, with permission from The Company of Biologists).

requiring reasonable depth perception. These crabs also are quick to spot potentially predatory birds and take cover quickly in response to rapidly looming objects. Electrophysiological studies of the crab visual system support their capability for acute movement detection (see c­ hapter 19 in this volume). Vigorous spike responses from movement-detecting neurons in the lobula of the semiterrestrial crab Neohelice granulata (= Chasmagnathus granulatus) have been obtained following stimulation of the contralateral, ipsilateral, and, simultaneously, both compound eyes (Sztarker and Tomsic 2004). These studies are backed up by neuroanatomical investigation of the lobula neuropil in Neohelice, revealing a columnar organization whose spacing corresponds closely with the retinotopic organization of the lamina, and characterized by the presence of prominent tangential interneurons, which, as in possibly homologous neurons in the insect lobula plate, respond to movement across the retina (Sztarker et al. 2005). It is thus clear that vision



Sensory Systems of Crustaceans

plays a vitally important role in the life of stomatopods and some species of brachyuran crabs; the extent to which this is true of most other crustacean taxa remains to be determined.

GEOMAGNETIC AND ELECTRICAL SENSES Th o u g h t h e y e s t y waves confound and swallow navigation up. . . Be bloody, bold and resolute. —William Shakespeare, Macbeth, iv. i.

Geomagnetic Sense There is compelling behavioral evidence that spiny lobsters possess not only a magnetic compass sense but also a positional map sense, both capabilities dependent on sensing the Earth’s magnetic field (see ­chapter 12 in this volume). Spiny lobsters forage at night, moving long distances from their daytime dens even on moonless nights, and returning before dawn. They are capable of returning to their home locations even when displaced by distances of up to five miles (Creaser and Travis 1950). Lobsters captured at a site in the Florida Keys and subjected to geomagnetic cues corresponding to sites either to the north or to the south of their home location will orient in an arena predominantly in a direction that would lead them back toward their home from their perceived locational displacement (Boles and Lohmann 2003, Lohmann 2010). Furthermore, lobsters that have actually been displaced either north or south from their home territory in the Florida Keys by several kilometers will subsequently orient in a direction that points toward their home site when blindfolded and tested in a large circular arena. A well-known phenomenon is the mass migration of P. argus from shallow waters to deeper environments during the fall and early winter. Along the west coast of Bimini in the Bahamas, the major direction of these migrations is from north to south, possibly using geomagnetic field alignment cues. The sense organs involved in the determination of geomagnetic cues in spiny lobsters have not been identified. Two major mechanistic hypotheses, one involving the alignment of microscopic, single domain magnetite crystals, and the other involving the intervention of radical pair reactions, have been proposed to explain the geomagnetic capabilities of birds, honeybees, and other animals. While there is significant evidence for the existence of biogenic magnetite crystalbearing cells in a number of animals, their specific mechanism of neuronal activation remains elusive. One model involves a direct mechanical linkage to ion channels (Lohmann 2010), but specific evidence is lacking. Even more hypothetical, but gaining some adherents (Solov’yov et al. 2010), is the possibility of radical pair reactions within photoreceptors in the eyes. According to this hypothesis, which involves the blue light-absorbing protein cryptochrome, the unpaired electron spins of the radical pairs precess around a local magnetic field that can interact with weak externally imposed magnetic (geomagnetic) fields. Resulting changes in electron transfer, involving both tryptophan and reduced flavin adenine dinucleotide, affect the extent of cryptochrome’s activation by light. However, whether any organisms, including spiny lobsters, use this mechanism to detect the Earth’s magnetic field is currently not known. Electrical Sensing Suggestions that aquatic crustaceans may be able to sense weak electrical currents in their environment have surfaced from time to time (Patullo and Macmillan 2007, 2010, Steullet et  al. 2007). There is now clear behavioral evidence from the crayfish Cherax destructor

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DeForest Mellon, Jr. and Cherax quadricarinatus that suddenly exposing these animals to environmental electric currents producing field strengths of 3–7 mV/cm arrests previously ongoing behavior, strongly suggesting that the animals are attending to the electric field generated by a dipole. Furthermore, these studies indicate that the effective fields are well within the amplitude and frequency range of those generated by muscle activity in swimming vertebrates and invertebrates (Fig. 3.6B), providing the possibility that they may be used by crayfish to detect nearby predators or prey (Patullo and Macmillan 2010). Steullet et al. (2007), using slightly different techniques and behavioral criteria in which the animals probed or grabbed at the electrodes generating a dipole field, found evidence that P.  clarkii can sense slowly (4–10 Hz) oscillating and/or DC electric fields of 20 mV/cm. As is the case with the geomagnetic sense, however, there is no clue concerning the sense organs responsible for this capability. In most cases, the high electrical resistance of crustacean cuticle must present a considerable barrier to the passage of electric current. Chemoreceptors, however, such as those found in bimodal sensilla discussed earlier in this chapter, could provide a potential low resistance transcuticular pathway, since their thin cuticle or access pore of necessity must be transparent to water soluble molecules at least as large as many amino acids and would present little resistance to the f low of ionic current. It has been pointed out (Steullet et  al. 2007)  that, while at least three crayfish species can detect electric fields at the high-threshold end of the sensitivity spectrum exhibited in aquatic vertebrates, all aquatic animals known to use electroreception for predation have specialized, well-defined electroreceptor organs. The lack of evidence of such organs on crayfish casts significant doubt that they normally use electroreception to locate prey.

MECHANORECEPTION B u t O for the touch of a vanished hand, And the sound of a voice that is still. —Alfred, Lord Tennyson, Break, Break, Break

Since the 1950s, mechanoreceptor neurons of decapod crustaceans have provided unique and fundamental insights to the basic understanding of cellular neurophysiology. The large size, interesting behaviors, and robust physiological tractability of decapods often has promoted them as subjects for neuroethological studies, and the large size of the neurons associated with many mechanoreceptors early on targeted them as uniquely appropriate subjects for electrophysiological examination. Primary—now classical—examples are the studies with multipolar abdominal stretch receptor neurons in lobsters and crayfish, which were crucial to our developing general understanding of the electrical events leading to impulse generation in all sensory cells. They also provided early insights into the mechanisms of synaptic inhibition (Wiersma et  al. 1953, Kuffler 1954, Eyzaguirre and Kuffler 1955a, 1955b, Kuffler and Eyzaguirre 1955, Edwards and Ottoson 1958, Erxleben 1989). Studies of other mechanoreceptor neurons in decapods have introduced entirely new concepts of receptor neuron physiology (Bush and Roberts 1968, Ripley et al. 1968) and have enlarged our understanding of impulse initiation and its relationship to cellular geometry (Mendelson 1963, 1966, Mellon and Kennedy 1964, Mellon and Kaars 1974). Finally, reflexive behaviors in crustaceans triggered or driven by mechanoreceptor afferents—both extero- and proprioceptors—are among the most completely understood of any animal in terms of their neural substrates (Fields and Kennedy 1965, Fields et al. 1967,



Sensory Systems of Crustaceans

Krasne 1969, Zucker et al. 1971, Clarac and Dando 1973, Clarac et al. 1978, Wine and Krasne 1982, Edwards et al. 1998, Herberholz et al. 2002). Because the crustacean cuticle serves as a skeleton, as with other arthropods, mechanoreception in crustaceans is primarily carried out through cuticular modifications, that is, through the elaboration of transcuticular sensilla, by means of muscular or chordotonal associations with the endocuticle or exoskeletal apodemes, or through direct innervation of regions of hypodermis beneath areas of soft cuticle. In these instances, movement of a sensillum or appendage joint generates strain to a region where sensory neuron dendrites are closely associated with or tied to the movable structure by means of connective tissue strands. In other instances, as in muscle receptor organs of the abdominal segments or the thoracicocoxopodite limb joint muscle receptors, the dendrites are embedded in regions of connective tissue intimately insinuated within the muscle itself. As alluded to earlier in this chapter, bimodal chemo-mechanoreceptor sensilla abound in crustaceans and, along with the purely mechanoreceptor sensilla, exhibit tremendous structural, interspecific diversity. In this section, I will examine some features of the exteroceptor and proprioceptor sensors among decapod crustaceans, attempting to correlate their features, placement, and structure with behavior where the connection is understood. Exteroceptors In most malacostracans, no surface or appendage, with the exception of the compound eyes, leg joint membranes, and abdominal soft intertergal cuticle is free of some form of mechanosensitive setae, and there is a striking diversity of sensillar forms among different species. Among the exclusively mechanoreceptive sensilla, two major categories can be recognized: hydrodynamic (near-field) receptors and contact mechanoreceptors. Hydrodynamic receptors are found on the antennules, cephalothorax, chelae, abdominal tergites, telson, and uropods. The first types to be described electrophysiologically were the hair-peg and hair-fan receptors on the lobster Homarus vulgaris (= H. gammarus) (Laverack 1962a, 1962b, 1963). These organs are found in shallow round pits on the chelae and carapace of lobsters in reasonably high density (25 cm-2). A round or oval plate at the bottom of the pit serves as the origin of a group of setae that projects up and slightly out of the pit, thereby being exposed to water movement (Fig. 3.7E). Both hair-peg organs, in which the setae surround a central peg, and hair-fan organs, which lack a peg, are sensitive to water movement. Hair-peg organs (Laverack 1962a) respond during their exposure to a continuous water current but are silent in the absence of coherent (advective) water currents. Hair-fan organs are more sensitive and respond to alternating pressure waves within the water up to sinusoidal frequencies of 80 Hz. They are dually innervated and certainly respond to stimuli within the acoustic near-field, and it has been suggested that they could possibly act as acoustic far-field detectors (Laverack 1962b) for stimuli at suitably low frequencies. There is no evidence, however, that any crustaceans respond behaviorally to sounds transmitted within the water column. In the crayfish P. clarkii, near-field hydrodynamic receptors occur on both lateral and medial flagella of the antennules (Mellon and Christison-Lagay 2008), branchiostegites (Mellon 1963), and dorsal surface of the telson (Wiese 1976). The morphology and innervation of some of the sensilla at these diverse locations is identical, and it is highly probable that they represent the same sensillum type (see Fig. 3.7A–d). In terms of their morphology, near-field receptors in P. clarkii are plumose (feathered) sensilla 100–150 μm in length and about 10 μm in diameter at their base. They are characterized by a planar array of ribbon-like filaments along the sensillum shaft, which tend to be shorter toward the distal end of the sensillar shaft (Fig. 3.7A,b1,c). The sensillar base itself tapers abruptly as it enters its cuticular socket, and it appears to be embedded in wrinkled, soft cuticle, allowing it to pivot freely within the socket. The base of the shaft

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Fig. 3.7. Crustacean hydrodynamic receptors:  SEM images. (A)  Standing feathered sensillum from the medial antennular flagellum of the crayfish Procambarus clarkii. The planar array of filaments is conspicuous. (B1) Higher magnification of the ribbon-like filaments from an antennular feathered sensillum. (B2) Base of an antennular feathered sensillum showing the prominent constriction at the socket collar. (C) “Pit-hair” hydrodynamic sensillum (arrow) on the branchiostegite of P. clarkii of a type described by Mellon (1963). The white asterisk indicates a “companion” seta, believed not to be innervated. (D)  Constricted base and socket collar of the sensillum shown in C. (E) Hair-fan organ from the cheliped of Homarus (from Laverack 1963, with permission from Elsevier). Scale bars: A, 50 μm; B1, 15 μm; B2, 8 μm; C, 50 μm; D, 10 μm; E, 40 μm.

forms a linear connection with the soft socket cuticle that serves as a hinge for the sensillum and appears to be the basis for its directional compliance (Mellon 2012). Feathered sensilla of types at all of the locations listed above are dually innervated (Mellon 1963, Wiese 1976, Mellon and Christison-Lagay 2008). Their sensory neurons are highly sensitive to angular displacements of the sensillum from its resting posture, especially in the preferred plane (threshold = 0.02 μm at 10 Hz for the tips of antennular feathered sensilla, but, of course, much less at the nexus with the sensory dendrite). Analysis of the spiking responses in the feathered sensilla of both the telson and the antennules indicates that frequency increases monotonically with increases in angular velocity (Wiese 1976, Mellon and Christison-Lagay 2008). Plumose sensilla sensitive to near-field vibrations have also been described on the claws of the crayfish C. destructor (Tautz and Sandeman 1980). Response thresholds of these sensilla at sinusoidally generated vibrations of 150 Hz, their best frequency, were close to 0.2 μm particle displacement amplitude, and response thresholds for feathered sensilla on the telson at 10–40 Hz were 0.1 μm—in both cases about an order of magnitude higher than described for the feathered sensilla on the crayfish antennules (Mellon and Christison-Lagay 2008). Like the hair-fan organs of lobsters, therefore, crayfish feathered/plumose sensilla appear to be adequately suited



Sensory Systems of Crustaceans

for the detection of near-field water particle movements generated by the rapid approach of potential predators or prey, and they undoubtedly constitute an important component of the afferent limb of giant fiber-mediated escape responses (e.g., Edwards et  al. 1998). Plummer et al. (1986) identified three general classes of interneurons in the sixth abdominal ganglion of P. clarkii that responded to low-frequency, high-frequency, and broad-band vibration stimuli in the water column, presumably following excitation of the telson/uropods’ near-field receptors. On the lateral antennular flagellum of P. clarkii, stimulation of even a single standing feathered sensillum can evoke a flagellar flick response (Mellon and Abdul Hamid 2012). This behavior, like similar actions in other decapods, is believed to reduce or remove the boundary layer around individual aesthetasc sensilla, subsequently exposing them to novel water parcels that may contain traces of odorants (see ­chapter 10 in this volume).

Contact Sensilla Bimodal contact sensilla were discussed at length at the beginning of this chapter and will not be discussed further. Unimodal contact sensilla are found conspicuously on the second antennae. Taylor (1975) characterized mechanosensory inputs from the crayfish antenna and classified them according whether they responded to vibrational, directional, or positional movement. He found that the afferents from the antennae course to the tritocerebrum, where they terminate in stratified neuropil in a topographic, stimulus modality-dependent manner (Taylor 1975). Tazaki (1977) examined spiking responses from two types of mechanoreceptors on the antennal f lagellum of the lobster H. gammarus. One type was clearly a near-field receptor, highly sensitive to oscillatory and linear def lections, and dually innervated. The other type was longer and also responded to hydrodynamic as well as tactile stimulation. Sandeman (1989) studied the antennae of the crayfish C. destructor and identified four major types of setae along the f lagellum, at least two of which are innervated and sensitive to mechanical displacement. The antennae are important organs for tactile localization (Sandeman 1985, Sandeman and Varju 1988); they respond to hydrodynamic inputs by rapidly moving toward and making contact with objects moving in their vicinity (Breithaupt et al. 1995) and can detect not only the direction but also the shape of objects that they encounter (Sandeman 1985). Statocysts Statocysts are essentially organs of equilibrium and technically are proprioceptors. Nonetheless, as they are basically cuticular structures armed with batteries of cuticular sensilla, they will be included in the exteroceptor section. Erroneously described as organs of hearing by 19th-century biologists (e.g., Huxley 1896), the decapod statocysts were convincingly shown to be organs of gravitational detection through the famous experiment of Kreidl (1893) in which he supplied a freshly molted caridean shrimp (Palaemon) with iron filings, which the animal used to replenish the sand grains lost along with the shed cuticular lining of the statocysts. Subsequent exposure to a strong magnet permitted Kreidl to modify the normal resting posture of the shrimp in a predictable way. Later work by Schöne (1954) and Dijkgraaf (1956) explored the compensatory eyestalk and postural reflexes generated in part by statocyst inputs to the brain (reviewed in Cohen and Dijkgraaf 1961). The statocyst of macruran decapods, exemplified by that of H. americanus (Cohen 1955), is rather straightforward. A cuticular-lined, fluid-filled open cavity on the dorsal surface of the first (coxopodite) antennular segment contains a statolith composed of loosely aggregated sand grains. The statolith rests on the floor of the statocyst and is surrounded by

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DeForest Mellon, Jr. four rows of sensilla, generally referred to as hook hairs. The hook hairs are in contact with the statolith and act as position receptors, signaling movement and displacement of the statolith as the animal’s position in the pitch and roll axes changes. Horizontal rotation (yaw) is sensed by fluid inertia within the statocyst cavity that acts on another type of sensillum, the thread hairs. Elaboration of the statocyst into a more precise organ of equilibrium has occurred in the brachyurans. In this group, exemplified by the crab Scylla serrata (Sandeman and Okajima 1972), the statocyst has the form of two joined toroids positioned orthogonally to one another, and forming horizontal and vertical fluid-filled interconnected canals (Fig. 3.9D,E). Fluid motion within each of the two canals is sensed by thread hairs and can provide the crab with exceedingly accurate information concerning angular acceleration in every spatial plane. A statolith at the bottom of the vertical canal is surrounded by hook hairs and senses positional changes in the crab’s posture. Many crabs have excellent vision, and static as well as dynamic eyestalk stabilization reflexes, largely generated by statocyst input, are the most robust among the decapods. Proprioceptors Crustaceans are remarkably well endowed with proprioceptors that measure joint movement and position, apodeme tension, cuticular stress, muscle length, and its dynamic changes, and even stretch of the abdominal ventral nerve cord. Electrical responses from several types of decapod mechanoreceptor sensilla are illustrated in Fig. 3.8A–E. Muscle Receptor Organs Crustacean muscle receptor organs (MROs) were first described in the abdominal and thoracic segments (Alexandrowicz 1951, 1952) of macrurans and later in the thoracicocoxopodite joint (Alexandrowicz and Whitear 1957). The decapod abdominal MROs are perhaps best known among the various proprioceptors in terms of their electrophysiology and their involvement in behavioral actions. Stephen Kuffler and Carlos Eyzaguirre immortalized the abdominal MRO sensory neurons in reptantians in their classic papers on excitation and inhibition in these cells, among the first electrophysiological studies that elucidated the nature of the receptor potential and the ionic basis for synaptic inhibition (Eyzaguirre and Kuffler 1955a, 1955b, Kuffler and Eyzaguirre 1955) A later study by Erxleben (1989) was among the first to demonstrate single-channel currents in the dendrites of MRO sensory neurons in response to membrane tension (Fig. 3.9C). In crayfish, shrimp, and lobsters, the abdominal MROs are activated by abdominal flexion, which passively stretches the associated muscle fibers, or by activation of their motor supply, which increases their isometric tension. Each abdominal hemisegment has an MRO consisting of a single pair of muscle fibers, one rapidly contracting (phasic) and supplied by a rapidly adapting sensory neuron, and one slowly contracting (tonic) fiber supplied by a slowly adapting sensory neuron (Fig. 3.9A). The MROs are situated in parallel with the superficial extensor muscles, and the “slow” MRO is involved in local regulation of abdominal postural control. In Homarus, the dendrites of its slowly adapting sensory neuron are embedded in a connective tissue matrix roughly halfway along the length of the muscle fiber and are sensitive to tension generated by abdominal flexion or by contraction of the muscle fiber via a motor neuron that it shares with the fibers of the superficial extensor muscle for that segment. In crayfish, there is no intercalated region of connective tissue in the muscle fiber. Excessive tension in the MRO due to any obstruction of intersegment shortening by the superficial extensor muscle is sensed by the slowly adapting sensory neuron, and its action potentials directly activate a second motor neuron that solely supplies the superficial extensor of that segment. The additional muscular activity can be sufficient to overcome the difficulty in segment shortening,

Fig. 3.8. Crustacean mechanoreceptors. (A) Responses from the paired axons of a standing feathered sensillum to sinusoidal movements of a stimulus probe at a frequency of 0.5 Hz. Extracellular records (from Mellon and Christison-Lagay 2008, with permission from the National Academy of Sciences). (B) Spike responses of paired neurons associated with near-field receptive sensillum on the telson of Procambarus clarkii in response to sinusoidal movement at 0.5 Hz (modified from Wiese 1976 with permission from American Physiological Society). (C1) and (C2) Spiking responses of neurons associated with a hair-fan organ on the cheliped of Homarus americanus to, respectively, sinusoidal movements at 2 Hz and 4 Hz (modified from Laverack 1963, with permission from Elsevier). (D) Single-channel currents from a primary dendrite of a slowly adapting MRO sensory neuron of the crayfish Orconectes limosus in response to slight pressure. Current amplitudes are shown at several different voltage displacements (holding potential, HP) from the resting membrane potential of the cell (modified from Erxleben 1989, with permission from Rockefeller University Press). (E1) Extracellular recording from receptor nerve (top trace), intracellular recording from S fiber of the crab thoracicocoxopodite MRO (middle trace), and in situ muscle stretch monitor (bottom trace) showing nonspiking response to stimulation. (E2) Graded variations in amplitude of the electrical response of a D fiber in the MRO (upper traces) to graded intensities of stretch. (E3) Responses of a T fiber in the crab thoracicocoxal MRO to graded velocities of stretch (lower traces). Calibrations: 100 μV extracellular and 20 mV intracellular. (E1 from Ripley et  al. 1968, with permission from Macmillan Publishers, E2 and E3 from Bush and Roberts 1971, with permission from The Company of Biologists).

A

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Thick accessory nerve Thin accessory nerve

Telson 6 5 4 3 2 1 Abdominal segments 5 4 Legs

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Mxpd 3 3 2 Cheliped Depressor receptor D fiber S fiber

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Fig. 3.9. Crustacean mechanoreceptors. (A)  Abdominal MRO of the crayfish indicating the paired receptor muscle fibers with their respective sensory neurons, motor axons, and inhibitory nerve fibers (from Alexandrowicz 1967, with permission from John Wiley & Sons). (B) Disposition of crayfish MROs in the abdominal (arrows) and thoracic segments (right side only; modified from Bush and Laverack 1982, with permission from Academic Press). (C) Thoracicocoxal MRO of the crab Carcinus maenas, indicating the receptor muscle and four receptor nerve fibers (from Bush and Roberts, 1971, with permission from The Company of Biologists). (D) Fused, orthogonal toroid model of the statocyst of the crab Scylla serrata, shown in E. Direction of fluid motion within the organ is shown for counter-clockwise yaw rotations. The left-hand model employs precise circular toroids, whereas the right-hand model more closely resembles the actual crab structure. (E) Diagram of the right-hand statocyst of Scylla, indicating the location of thread hairs, free hook hairs, and the statolith organ. Open arrows indicate the flow of fluid across and around the sensory cushion in response to rotation of the crab in a counterclockwise direction (D and E from Sandeman and Okajima 1972, with permission from The Company of Biologists). (F)  Pereopod of the lobster Homarus americanus indicating the position of (arrowheads) chordotonal organs and other proprioceptors at the limb joints. Inset: detail of the chordotonal organ at the propodite-dactylopodite joint in the crab Carcinus maenas. Detail of the organ is shown at higher magnification beneath the diagram of the joint. Abbreviations: d, dactylopodite; N, limb nerve; n, nerve strand from chordotonal organ; o, chordotonal organ embedded in a connective tissue strand; p, propodite, (*) origin of the connective tissue strand on the dactylopodite; (§) insertion of connective tissue strand on the dactyl flexor muscle apodeme (F from Clarac 1977, with permission from Plenum Press; inset from Burke 1954, with permission from The Company of Biologists).



Sensory Systems of Crustaceans

thereby releasing the tension in the MRO and reducing the output of its sensory neuron. The slowly adapting crayfish MROs and their motor supply thereby act as length-tension servos, which locally regulate actions generated through central commands (Fields and Kennedy 1965, Fields et al. 1967). The rapidly adapting neurons of the “fast” MROs in the abdominal segments are critical elements in the crayfish tail-flip escape reflexes. Although command-derived inhibition of these sensory neurons is mediated by the lateral giant action potential at the same time that it excites the fast flexor motor neurons, the subsequent powerful flexion produced during a tail flip maximally excites the rapidly adapting neurons associated with the phasic MROs of the abdominal segments, and their high-frequency impulse outputs act centrally to directly excite motor neurons to the fast extensor muscle fibers, reextending the abdomen following the initial flexion. This action prepares the animals for subsequent escape swimming or a medial giant fiber-mediated tail flip if required (Wine and Krasne 1982). The thoracicocoxal MRO of brachyurans discovered by Alexandrowicz and Whitear (1957) and later examined electrophysiologically by Bush and coworkers (reviewed by Bush 1977), like the chordotonal organs discussed below, mediate resistance reflexes that control and stiffen joint movement at the base of the leg. At least one of the sensory neurons associated with this MRO excites spiking activity in the motor neurons of the coxopodite promoter muscle, which lies in parallel to the MRO across the thoracicocoxal joint. Under isometric conditions, stimulation of the promoter motor neurons and subsequent contraction of the promoter muscle also generates reflex firing in the promoter. This positive feedback may act as an assistance in moving the limb forward under some circumstances (see also Evoy and Cohen 1971, Barnes et al. 1972, Evoy and Fourtner 1973, Bush et al. 1975, Clarac and Vedel 1975, Kennedy and Davis 1977). In brachyuran crabs, the three major sensory neurons associated with these organs conduct receptor potentials to their neuronal targets in the thoracic ganglia by passive electrotonic spread along their unusually large-diameter dendrites, without the intervention of action potentials. They are also unusual among arthropod sensory neurons in that their cell bodies reside within central ganglia. Nonspiking mechanoreceptor afferents also occur in the uropods of the sand crab Emerita (Paul 1972). Like the sensory neurons associated with the thoracicocoxal organ, the cell bodies of these neurons reside in the CNS, and they also appear to generate resistance reflexes in uropod power stroke muscles. The existence of nonspiking mechanoreceptor neurons raises questions about the properties of the sensory neuron dendrites that promote decremental conduction of the receptor potential and about the dynamics of synaptic transmitter release machinery at the dendritic targets in the CNS. Furthermore, sensory neurons subserving other limb joint receptors in crabs and other decapods employ conventional impulse conduction mechanisms, although admittedly they are much further away from their central targets, making decremental conduction highly problematical. Is there a unique behavioral importance to the thoracicocoxal joint and its promoter muscle that demands the more subtle control obtained by continuous, rather than impulsive, transmitter release from the MRO afferent terminals? Chordotonal (Scolopidial) Organs In typical decapods, proprioception at other joints in the walking limbs, other thoracic appendages, antennae, and antennules is carried out primarily by chordotonal organs, connective tissue sheets or strands into which are inserted the dendrites of (usually) multiple bipolar mechanosensory neurons (reviewed by Clarac 1977, and Bush and Laverack 1982). In general these organs sense joint position and movement (both extension and flexion). Subtypes of chordotonal organs are recognized, depending on whether they are associated with connective tissue

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DeForest Mellon, Jr. strands, sheets, or in a few instances, directly with muscle, such as the accessory flexor muscle at the ischiomeropodite (I-M) joint in Reptantia (Mill 1976). Chordotonal organs have also been referred to as scolopidial organs due to the presence of a spindle-shaped, electron-dense structure, the scolopale, which surrounds a portion of the distal dendritic segments of associated sensory neurons and ties them to the connective tissue into which they are embedded. It is believed (Lowe et al. 1973, Mill and Lowe 1973) that the adequate stimulus for chordotonal organ activation is stretch of the dendritic segments within the scolopales. The behavioral role of chordotonal organs is of considerable interest. While they undoubtedly mediate resistance reflexes that can oppose the movement that excites them (Barnes et al. 1972, Evoy and Fourtner 1973, Clarac 1977, 1981), they are also believed to be involved during voluntary movements, as are the thoracicocoxal MROs mentioned above, in assisting appendage movements through positive feedback (Clarac and Vedel 1975, Bush 1977, Kennedy and Davis 1977, Vedel 1980). This is especially true in the Brachyura, in which the evolution of limb control appears to have proceeded at a pace far more advanced than that found in the Astacida, Palinura, or Homarida. Anyone who has ever watched the impressive locomotory performance of ocypodid or portunid crabs, for example, cannot help but be impressed with their speed and coordination. By primate analogies, if crayfish and lobsters are the tarsiers and lorises of the crustacean world, crabs are surely the lemurs! Types of proprioceptor termed myochordotonal organs have been described for the I-M joint in all decapods, but have been most extensively studied in reptantian species, as reviewed by Mill (1976) and Bush and Laverack (1982). There are actually three separate organs at the I-M joint. IM1 originates on the meropodite reductor muscle; IMa is attached to the accessory flexor muscle, and it is activated by flexion of the merocarpopodite joint, brought about either by active or passive shortening of the accessory flexor muscle. A third myochordotonal organ, sometimes referred to as Barth’s organ, is stretched by passive merocarpopodite joint extension or contraction of the accessory flexor muscle. Chordotonal organs are thus of overarching importance in the control of the decapod pereopods but are also found (in significantly reduced numbers) in the maxillipeds and first and second antennae (Mill 1976). Several other types of proprioceptors have been described in crustaceans and have been reviewed by Bush and Laverack (1982). They include innervated elastic sheets and strands, which differ from chordotonal organs in lacking a scolopale, and in which the dendrites of associated unitary or multipolar sensory neurons are directly embedded and attached to elastic connective tissue (Mill 1976). Of special note are the nerve cord stretch receptors initially discovered by Hughes and Wiersma (1960) in Procambarus and examined more extensively by Grobstein (1973). They include both phasic and tonic units that are active when the crayfish, respectively, reextends its abdomen or is in a resting posture with the abdomen extended. The dendrites of the sensory neurons involved probably are embedded in the sheath surrounding the abdominal ventral nerve cord. Cuticular strain or stress receptors are found associated with regions of soft cuticle at the base of the pereopods in decapods, adjacent to the preformed autotomy plane between the basipodite and ischiopodite segments. They resemble chordotonal organs in that the dendrites of multiple bipolar sensory neurons insert into scolopales that are themselves embedded in the cuticle. Areas of the hypodermis beneath the soft cuticle of the intersternal abdominal segments in the crayfish P. clarkii are locally supplied by end bulb terminals of bi- and tripolar sensory neurons having somata in abdominal ganglionic nerve roots 1 and 2 (Pabst and Kennedy 1967). Unusually for a crustacean, the cell bodies are located close to their ganglion of entry, and their distal dendrites, which are quite long and transmit action potentials, may branch a number of times to supply several areas of soft cuticle that can be some distance apart. These regions occur



Sensory Systems of Crustaceans

primarily near the bases of the swimmerets and, especially, along the points of insertion of the superficial flexor muscles. Spiking activity in these hypodermal receptors generated by punctate depression of the soft cuticle is accompanied by inhibition of tonic activity in the motor neurons to the superficial flexors. The question of whether crustaceans are sensitive to stimuli in the acoustic far-field is raised by the fact that a number of decapods generate sounds that are audible to humans. Snapping shrimp (Caridea, Alpheidae), of course, snap (Versluis et al. 2000); Panulirus stridulates (Patek 2001, 2002); mantis shrimp rumble (Patek and Caldwell 2006); crabs can creak (Field et  al. 1987); and Homarus contracts its antennal remoter muscle at over 100 Hz to generate a buzz that can be felt when grasping the antennal shaft (Mendelson 1969). Most of these sounds (with the possible exception of that made by snapping shrimps) are believed to be antipredator cues (Patek et al. 2009). However, to date, in the absence of anatomical evidence for a possible organ of hearing in any crustacean, and with no credible behavioral data to suggest that auditory communication between conspecifics occurs, it must be concluded that no crustacean can sense fluid particle vibrations in the acoustic far-field. Some behavioral evidence for hydrostatic pressure sensitivity in benthic and planktonic crustaceans has been previously documented (Knight-Jones and Qasim 1955, Knight-Jones and Morgan 1966, Naylor and Atkinson 1972), but until recently there has been no physiologic evidence for a crustacean hydrostatic sensor. Now, however, reports have emerged that sensilla within the statocyst organ of crabs can act as hydrostatic pressure sensors (Fraser and MacDonald 1994). Thread hairs within the statocyst have fluid-filled lymph cavities surrounding the chorda-like attachments of the associated sensory neuron dendrites to the hair shaft. Although fluid motion within the statocyst canals (Fig. 3.9E) normally constitutes the adequate stimulus for these sensilla, small modifications in lymph cavity fluid volume occasioned by hydrostatic pressure (depth) changes are believed to act via a piston model, slightly displacing the chorda and effecting reduction in dendritic membrane potential thereby. More recent studies showed that simultaneous changes in pressure and temperature affected spiking frequency during rotation of the crab, findings that are consistent with the piston model (O’Callaghan and Fraser 2010). The thread hair piston model represents the sole hydrostatic pressure detection mechanism hypothesized for crustaceans, an idea that had previously not received much attention due to the lack of known gas-filled structures that could obviously serve as a hydrostatic detection system. While water is far less compressible than gas at comparable temperatures, it is not infinitely so, and a one-bar pressure change could change the length of the fluid volume inside a statocyst thread hair by approximately 0.02 μm (Fraser and MacDonald 1994). The piston model is supported by the fact that this length change, if transmitted undiminished to the chorda within the hair by piston-like action, would move the dendrite attachment site by an amount equal to or greater than threshold movements reported for near-field sensilla in the crayfish (Wiese 1976, Mellon and Christison-Lagay 2008).

FUTURE DIRECTIONS Not all aspects of peripheral sense organ physiology have been successfully attacked at this time; aspects of signal transduction cascades in chemoreception and photoreception, for example, remain of significant interest to those examining sensory systems at the molecular level. But, inevitably, as questions concerning sense organ physiology per se are elucidated, the attention of those interested in crustacean sensory systems will be directed increasingly to the CNS. Problems of higher order sensory operations within the crustacean brain have barely been touched, no doubt owing in great part to problems (not normally encountered in insects) with

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DeForest Mellon, Jr. keeping the brain alive and functioning at an appropriate physiological level. These technical problems can now be overcome in at least larger crustaceans by brain perfusion or in vivo recording methods, which will be necessary to address questions of multimodal integration, memory and learning, and the modification of locomotory activity by higher level centers.

CONCLUSIONS The most primitive marine crustaceans first appear in the fossil records in the late Cambrian Period; since that time they have radiated into a highly successful group of aquatic organisms with a few members even invading the terrestrial habitat. During their evolution, crustaceans have acquired some highly sophisticated sensory systems, some of them rivaling or even exceeding the capabilities of those found among advanced vertebrates. The impressive statocyst and proprioceptive regulation of limb control in brachyurans comes to mind, as does the visual system of stomatopods. The olfactory systems found in crustaceans can be extremely sensitive and, if neuroanatomy is any guide, have more potential for spectral complexity than those found among insects. In crustaceans, moreover, the olfactory system can be critically important in intraspecific communication. Furthermore, recent findings concerning neurogenesis in crustacean olfactory systems offer the possibility that their capabilities can expand throughout life spans that in some instances encompass decades. The crustacean olfactory system clearly offers significant opportunities for examining not only basic cellular properties involved with odor perception but also the mechanisms through which novel odors are incorporated into a cellular memory. On the other hand, the lack of evidence strongly suggests that crustaceans cannot hear sounds. This sensory modality, which is so critically important among many insect groups, appears not to have developed in the evolutionary history of aquatic arthropods. In vertebrates, organs of hearing are derived from sensory structures (neuromasts) that originally served as near-field receptors and later were modified to detect energies in the acoustic far-field. We can only speculate on the conditions that prevented a similar transformation in the Crustacea of near-field receptors, with which they are so well endowed. For sixty years, crustacean sensory systems have been instrumental in shaping our understanding of basic physiological mechanisms in sense organs across modalities. These mines are far from exhausted. Furthermore, as pointed out by Bush and Laverack (1982), with the notable exception of stomatopods, the majority of this work has emerged from experimental work with decapods, and “The remainder of the Crustacea have barely been touched.”

ACKNOWLEDGMENTS The author is indebted to Ms. Jan Redick for her skill and efforts in producing the image of Fig. 3.1C.

REFERENCES Alexander, C.G. 1977. Antennal sense organs in the isopod Ligia oceanica. Marine Behaviour and Physiology 5:61–77. Alexandrowicz, J.S. 1951. Muscle receptor organs in the abdomen of Homarus vulgaris and Palinurus vulgaris. Quarterly Journal of Microscopical Science 92:163–199.



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Alexandrowicz, J.S. 1952. Receptor elements in the thoracic muscles of Homarus vulgaris and Palinurus vulgaris. Quarterly Journal of Microscopical Science 93:315–346. Alexandrowicz, J.S. 1967. Receptor organs in the thoracic and abdominal muscles of Crustacea. Biological Reviews of the Cambridge Philosophical Society 42:288–326. Alexandrowicz, J.S., and M. Whitear. 1957. Receptor elements in the coxal region of Decapoda Crustacea. Journal of the Marine Biological Association of the United Kingdom 36:603–628. Altner, I., H. Hatt, and H. Altner. 1983. Structural properties of bimodal chemo- and mechanosensitive setae on the pereopod chelae of the crayfish, Austropotamobius torrentium. Cell and Tissue Research 228:357–374. Anderson, P.A.V., and B.W. Ache. 1985. Voltage- and current-clamp recordings of the receptor potential in olfactory receptor cells in situ. Brain Research 338:273–280. Ball, E.E., and A.N. Cowan. 1977. Ultrastructure of the antennal sensilla of Acetes (Crustacea, Decapoda, Natantia, Sergestidae). Philosophical Transactions of the Royal Society of London B 277:429–457. Barnes, W.J.P., C.P. Spirito, and W.H. Evoy. 1972. Nervous control of walking in the crab, Cardiosoma guanhumi. II. Role of resistance reflexes in walking. Zeitschrift für vergleichende Physiologie 76:16–31. Berry, F.C., and T. Breithaupt. 2010. To signal or not to signal? Chemical communication by urine-borne signals mirrors sexual conflict in crayfish. BMC Biology 8:25. Bethe, A. 1897. Das Nervensystem von Carcinus maenas. Archive für mikroskopische Anatomie und Entwicklungsmechanik 50:460–546. Bobkov, Y.V., and B.W. Ache. 2007. Intrinsically bursting olfactory receptor neurons. Journal of Neurophysiology 97:1052–1057. Bobkov, Y.V., A. Pezier, E.A. Corey, and B.W. Ache. 2010. Phosphotidylinositol 4,5bisphosphate-dependent regulation of the output in lobster olfactory receptor neurons. Journal of Experimental Biology 213:1417–1424. Boles, L.C., and K.J. Lohmann. 2003. True navigation and magnetic maps in spiny lobsters. Nature 421:60–63. Breithaupt, T., and P. Eger. 2002. Urine makes the difference: chemical communication in fighting crayfish made visible. Journal of Experimental Biology 205:1221–1231. Breithaupt, T., B. Schmitz, and J. Tautz. 1995. Hydrodynamic orientation of crayfish (Procambarus clarkii) to swimming fish prey. Journal of Comparative Physiology A 177:481–491. Burke, W. 1954. An organ for proprioception and vibration sense in Carcinus maenas. Journal of Experimental Biology 31:127–137. Bush, B.M.H. 1977. Non-impulsive afferent coding and stretch reflexes in crabs. Pages 439–460 in G. Hoyle, editor. Identified neurons and behavior of arthropods. Plenum, New York. Bush, B.M.H., and M.S. Laverack. 1982. Mechanoreception. Pages 399–468 in H.L. Atwood and D.C. Sandeman, editors. The biology of Crustacea, Vol. 3, Neurobiology: structure and function. Academic Press, New York. Bush, B.M.H., and A. Roberts. 1968. Resistance reflexes from a crab muscle receptor without impulses. Nature 218:1171–1173. Bush, B.M.H., and A. Roberts. 1971. Coxal muscle receptors in the crab: the receptor potentials of S and T fibers in response to ramp stretches. Journal of Experimental Biology 55:813–832. Bush, B.M.H., D.H. Godden, and G.A. MacDonald. 1975. Voltage clamping of non-impulsive afferents of the crab thoracic-coxal muscle receptor. Journal of Physiology 245:3–5. Cate, H.S., and C.D. Derby. 2002. Ultrastructure and physiology of the hooded sensillum, a bimodal chemo-mechanosensillum of lobsters. Journal of Comparative Neurology 442:293–307. Chiou, T.-H., S. Kleinlogel, T. Cronin, R. Caldwell, B. Loeffler, A. Siddiqi, A. Goldizen, and J. Marshall. 2008. Circular polarization vision in a stomatopod crustacean. Current Biology 18:429–434. Clarac, F. 1977. Motor coordination in crustacean limbs. Pages 167–186 in G. Hoyle, editor. Identified neurons and behavior of arthropods. Plenum, New York. Clarac, F. 1981. Postural reflexes coordinating walking in a rock lobster. Journal of Experimental Biology 90:333–337.

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78

DeForest Mellon, Jr. Clarac, F., and M.R. Dando. 1973. Tension receptor reflexes in the walking leg of the crab, Cancer pagurus. Nature 243:94–95. Clarac, F., and J.P. Vedel. 1975. Proprioception by chordotonal and myochordotonal organs in the walking legs of the rock lobster, Palinurus vulgaris. Marine Behaviour and Physiology 3:157–165. Clarac, F., J.P. Vedel, and B.M.H. Bush. 1978. Intersegmental coordination by a single joint receptor organ (CB) in rock lobsters walking legs. Journal of Experimental Biology 73:29–46. Cohen, M.J. 1955. The function of receptors in the statocyst of the lobster Homarus americanus. Journal of Physiology 130:9–34. Cohen, M.J., and S. Dijkgraaf. 1961. Mechanoreception. Pages 65–108 in T.H. Waterman, editor. The physiology of Crustacea, Vol. 2, Sense organs, integration and behavior. Academic Press, New York. Creaser, E.P., and D. Travis. 1950. Evidence of a homing instinct in the Bermuda spiny lobster. Science 112:169–170. Cronin, T.W., N.J. Marshall, and R.L. Caldwell. 1994. The intrarhabdomal filters in the retinas of mantis shrimps. Vision Research 34:274–291. Cronin, T.W., N.J. Marshall, N.J., and R.L. Caldwell. 2000. Spectral tuning and the visual ecology of mantis shrimps. Philosophical Transactions of the Royal Society London B 355:1263–1267. Debelius, H. 1999. Crustacea guide of the world. Ikan, Frankfurt. Derby, C.D. 1989. Physiology of sensory neurons in morphologically identified cuticular sensilla of crustaceans. Pages 27–47 in B. Felgenhauer, L. Watling, and A.B. Thistle, editors. Functional morphology of feeding and grooming in Crustacea. A.A. Balkema, Rotterdam. Derby, C.D., and J. Atema. 1982. The function of chemo- and mechanoreceptors in lobster (Homarus americanus) feeding behaviour. Journal of Experimental Biology 98:317–327. Detto, T., P.R. Backwell, J.M. Hemmi, and J. Zeil. 2006. Visually mediated species and neighbour recognition in fiddler crabs (Uca mjoebergi and Uca capricornis). Proceedings of the Royal Society of London Series B: Biological Sciences 273:1661–1666. Devine, D.V., and J. Atema. 1982. Function of chemoreceptor organs in spatial orientation of the lobster, Homarus americanus: differences and overlap. Biological Bulletin 163:144–153. Dijkgraaf, S. 1956. The structure and function of the statocyst in crabs. Experientia 12:394–396. Edwards, C., and D. Ottoson. 1958. The site of impulse initiation in a nerve cell of a crustacean stretch receptor. Journal of Physiology, London 143:138–148. Edwards, D.H. 1984. Crayfish extraretinal photoreception. I. Behavioral and motoneuronal responses to abdominal illumination. Journal of Experimental Biology 109:291–306. Edwards, D.H., S.R. Yeh, and F.B. Krasne. 1998. Neuronal coincidence detection by voltage-sensitive electrical synapses. Proceedings of the National Academy of Sciences of the United States of America 95:7145–7150. Eguchi, E. 1999. Polarized light vision and rhabdom. Pages 33–46 in E. Eguchi and Y. Tominaga, editors. Atlas of arthropod sensory receptors. Springer, New York. Erxleben, C. 1989. Stretch-activated current through single ion channels in the abdominal stretch receptor organ of the crayfish. Journal of General Physiology 94:1071–1083. Evoy, W.H., and M.J. Cohen. 1971. Central and peripheral control of arthropod movements. Advances in Comparative Physiology and Biochemistry 4:225–266. Evoy, W.H., and C.R. Fourtner. 1973. Nervous control of walking in the crab Cardisoma guanhumi. III. Proprioceptive influences on intra- and intersegmental coordination. Journal of Comparative Physiology 83:303–318. Exner, S. 1891. Die Physiologie der facettierten Augen von Krebsen und Insekten. Deuticke, Vienna. Eyzaguirre, C., and S.W. Kuff ler. 1955a. Processes of excitation in the dendrites and in the soma of single isolated sensory nerve cells of the lobster and crayfish. Journal of General Physiology 39:87–119. Eyzaguirre, C., and S.W. Kuffler. 1955b. Further study of soma, dendrite, and axon excitation in single neurons. Journal of General Physiology 39:120–153. Fadool, D.A., and B.W. Ache. 1992. Plasma membrane inositol 1,4,5-trisphosphate-activated channels mediate signal transduction in lobster olfactory receptor neurons. Neuron 9:907–918. Fadool, D.A., W.C. Michel, and B.W. Ache. 1993. Odor sensitivity of cultured lobster olfactory receptor neurons is not dependent on process formation. Journal of Experimental Biology 174:215–233.



Sensory Systems of Crustaceans

Fahrenbach, W.H. 1964. The fine structure of a nauplius eye. Zeitschrift für Zellforschung und Mikroskopische Anatomie 62:182–197. Fay, R.R. 1973. Multisensory interactions in control of eye-stalk rotation response in the crayfish Procambarus clarkii. Journal of Comparative and Physiological Psychology 84:527–533. Fields, H.L., and D. Kennedy. 1965. Functional role of muscle receptor organs in crayfish. Nature 206:1232–1237. Fields, H.L., W.H. Evoy, and D. Kennedy. 1967. Reflex role played by efferent control of an invertebrate stretch receptor. Journal of Neurophysiology 30:859–874. Field, L.H., A. Evans, and D.L. Macmillan. 1987. Sound production and stridulatory structures in hermit crabs of the genus Trizopagurus. Journal of Marine Biological Association of the United Kingdom 67:89–110. Fraser, P.J., and A.G. MacDonald. 1994. Crab hydrostatic pressure sensors. Nature 371:383–384. Ghiradella, H.T., J.F. Case, and J. Cronshaw. 1968. Fine structure of the aesthetasc hairs of Coenobita compressus Edwards. Journal of Morphology 124:361–385. Gill, C.W., and C.J. Crisp. 1985. Sensitivity of intact and antennule amputated copepods to water disturbance. Marine Ecology Progress Series 21:221–227. Grasso, F.W., and J.A. Basil. 2002. How lobsters, crayfishes, and crabs locate sources of odor: current perspectives and future directions. Current Opinion in Neurobiology 12:721–727. Gray, E.G. 1960. The fine structure of an insect ear. Philosophical Transactions of the Royal Society London, B 243:75–94. Grobstein, P. 1973. Extension sensitivity in the crayfish abdomen. I. Neurons monitoring cord length. Journal of Comparative Physiology 86:331–348. Guse, G.-W. 1978. Antennal sensilla of Neomysis integer (Leach). Protoplasma 95:145–161. Hama, K. 1961. A photoreceptor-like structure in the ventral nerve cord of the crayfish, Cambarus virilus. Anatomical Record 140:329–336. Hansson, B., S. Harzsch, M. Knaden, and M. Stensmyr. 2011. The neural and behavioral basis of chemical communication in terrestrial crustaceans. Pages 149–173 in T. Breithaupt and M. Thiel, editors. Chemical communication in crustaceans. Springer, New York. Hatt, H., and B.W. Ache. 1994. Cyclic nucleotide- and inositol phosphate-gated ion channels in lobster olfactory receptor neurons. Proceedings of the National Academy of Sciences of the United States of America 91:6264–6268. Hemmi, J.M., and J. Zeil. 2003. Robust judgement of inter-object distance by an arthropod. Nature 421:160–163. Herberholz, J., B.L. Antonsen, and D.H. Edwards. 2002. A lateral excitatory network in the escape circuit of the crayfish. Journal of Neuroscience 22:9078–9085. Hughes, G.M., and C.A.G. Wiersma. 1960. Neuronal pathways and synaptic connections in the abdominal cord in the crayfish. Journal of Experimental Biology 37:291–307. Huxley, T.H. 1896. The crayfish. Kegan Paul, Trench, Trübner, London. Johnson, B.R., and J. Atema. 1983. Narrow-spectrum chemoreceptor cells in the antennules of the American lobster, Homarus americanus. Neuroscience Letters 41:145–150. Johnson, M.E., and J. Atema. 2005. The olfactory pathway for individual recognition in the American lobster Homarus americanus. Journal of Experimental Biology 208:2865–2872. Katona, S.K. 1973. Evidence for sex pheromones in planktonic copepods. Limnology and Oceanography 18:574–583. Kennedy, D. 1958. Electrical activity of a “primitive” photoreceptor. Annals of the New York Academy of Sciences 74:329–336. Kennedy, D. 1963. Physiology of photoreceptor neurons in the abdominal nerve cord of the crayfish. Journal of General Physiology 46:551–572. Kennedy, D., and W.J. Davis. 1977. The organization of invertebrate motor systems. Pages 1023–1087 in E.R. Kandel, editor. Handbook of physiology, Vol. 2, Neurophysiology. American Physiological Society. Bethesda, MD. Knight-Jones, E.W., and E. Morgan. 1966. Responses of marine animals to changes in hydrostatic pressure. Annual Review of Oceanography and Marine Biology 4:267–299.

79

80

DeForest Mellon, Jr. Knight-Jones, E.W., and S.Z. Qasim. 1955. Responses of some marine plankton animals to changes in hydrostatic pressure. Nature 175:941–942. Krasne, F.W. 1969. Excitation and habituation of the crayfish escape reflex: the depolarizing response in lateral giant fibers of the isolated abdomen. Journal of Experimental Biology 50:29–46. Kreidl, A. 1893. Weitere Beitrage zur Physiologie des Ohrlabyrinthes (II. Mitth.) Versuch an Krebsen. Sitzbergen Akademie Wissenschaften Wien Math-Naturwissenschaften Kl. Abteilen III 102:149–174. Kuffler, S.W. 1954. Mechanisms of activation and motor control of stretch receptors of lobsters and crayfish. Journal of Neurophysiology 17:558–574. Kuffler, S.W., and C. Eyzaguirre. 1955. Synaptic inhibition in an isolated nerve cell. Journal of Neurophysiology 39:155–184. Land, M.F. 1976. Superposition images are formed by reflection in the eyes of some oceanic decapod Crustacea. Nature 263:764–765. Land, M.F. 1980. Compound eyes: old and new optical mechanisms. Nature 287:681–686. Larimer, J.L. 1966. The presence of a functional caudal photoreceptor in blind cavernicolous crayfish. Nature 210:204–205. Larimer, J.L., D.L. Trevino, and E.A. Ashby. 1966. A comparison of spectral sensitivities of caudal photoreceptors of epigeal and cavernicolous crayfish. Comparative Biochemistry and Physiology 19:409–415. Laverack, M.S. 1962a. Responses of cuticular sense organs of the lobster, Homarus vulgaris (Crustacea). I. Hair-peg organs as water current receptors. Comparative Biochemistry and Physiology 5:319–325. Laverack, M.S. 1962b. Responses of cuticular sense organs of the lobster, Homarus vulgaris (Crustacea). II. Hair-fan organs as pressure receptors. Comparative Biochemistry and Physiology 6:137–145. Laverack, M.S. 1963. Responses of cuticular sense organs of the lobster, Homarus vulgaris (Crustacea). III. Activity invoked in sense organs of the carapace. Comparative Biochemistry and Physiology 10:261–272. Lenz, P.H., and J. Yen. 1993. Distal setal mechanoreceptors of the first antennae of marine copepods. Bulletin of Marine Science 53:170–179. Liden, W.H., and J. Herberholz. 2008. Behavioral and neural responses of juvenile crayfish to moving shadows. Journal of Experimental Biology 211:1355–1361. Lohmann, K.J. 2010. Magnetic-field perception. Nature 464:1140–1142. Lowe, D.A., P.A. Mill, and M.F. Knapp. 1973. The fine structure of the PD proprioceptor of Cancer pagurus. II. The position sensitive cells. Proceedings of the Royal Society of London, Series B: Biological Sciences 184:199–205. Marshall, N.J., T.W. Cronin, and N. Sashar. 1999. Behavioral evidence for polarization vision in stomatopods reveals a potential channel for communication. Current Biology 9:755–758. Martin, G., P.P. Jaros, J. Chaigneau, and V.B. Meyer-Rochow. 1995. Intracerebral ocelli in the giant Antarctic slater Glyptonotus antarcticus (Isopoda: Valvifera). Journal of Crustacean Biology 15:228–235. McClintock, T.S., and B.W. Ache. 1989. Histamine directly gates a chloride channel in lobster olfactory receptor neurons. Proceedings of the National Academy of Sciences of the United States of America 86:8137–8141. Mellon, DeF. Jr. 1963. Electrical responses from dually-innervated tactile receptors on the thorax of the crayfish. Journal of Experimental Biology 40:137–148. Mellon, DeF. Jr. 1997. Physiological characterization of antennular flicking reflexes in the crayfish. Journal of Comparative Physiology A 180:553–565. Mellon, DeF. Jr. 2000. Convergence of multimodal sensory input onto higher level neurons of the crayfish olfactory pathway. Journal of Neurophysiology 84:3043–3055. Mellon, DeF. Jr. 2005. Integration of hydrodynamic and odorant inputs by local interneurons of the crayfish deutocerebrum. Journal of Experimental Biology 208:3711–3720. Mellon, DeF. Jr. 2012. Smelling, feeling, tasting and touching: Behavioral and neural integration of chemosensory and mechanosensory antennular inputs in the crayfish. Journal of Experimental Biology 215:2163–2172.



Sensory Systems of Crustaceans

Mellon, DeF., and O. Abdul Hamid. 2012. Identified antennular near-field receptors trigger reflex flicking in the crayfish. Journal of Experimental Biology 215:1559–1566. Mellon, DeF., and V.E. Alones. 1995. Identification of three classes of multiglomerular, broad-spectrum neurons in the crayfish olfactory midbrain by correlated patterns of electrical activity and dendritic arborization. Journal of Comparative Physiology A 177:55–71. Mellon, DeF., and K. Christison-Lagay. 2008. A mechanism for neuronal coincidence revealed in the crayfish antennule. Proceedings of the National Academy of Sciences of the United States of America 105:14626–14631. Mellon, DeF. Jr., and C. Kaars. 1974. Role of regional cellular geometry in conduction of excitation along a sensory axon. Journal of Neurophysiology 37:1228–1238. Mellon, DeF. Jr., and D. Kennedy. 1964. Impulse origin and propagation in a bipolar sensory neuron. Journal of General Physiology 47:487–499. Mellon, DeF. Jr., and E.D. Lorton. 1977. Reflex actions of the functional divisions in the crayfish oculomotor system. Journal of Comparative Physiology A 121:367–380. Mellon, DeF. Jr., and S.D. Munger. 1990. Nontopographic projection of olfactory sensory neurons in the crayfish brain. Journal of Comparative Neurology 296:253–262. Mellon, DeF. Jr., H.R. Tuten, and J. Redick. 1989. Distribution of radioactive leucine following uptake by olfactory sensory neurons in normal and heteromorphic crayfish antennules. Journal of Comparative Neurology 280:645–662. Mendelson, M. 1963. Some factors in the activation of crab movement receptors. Journal of Experimental Biology 40:157–169. Mendelson, M. 1966. The site of impulse initiation in bipolar receptor neurons in Callinectes sapidus L. Journal of Experimental Biology 45:411–420. Mendelson, M. 1969. Electrical and mechanical characteristics of a very fast lobster muscle. Journal of Cell Biology 42:548–563. Mill, P.J. 1976. Chordotonal organs of crustacean appendages. Pages 243–298 in P.J. Mill, editor. Structure and function of proprioceptors in the invertebrates. Chapman-Hall, London. Mill, P.J., and D.A. Lowe. 1973. The fine structure of the PD proprioceptor of Cancer pagurus. I. The receptor strand and the movement sensitive cells. Proceedings of the Royal Society of London Series B: Biological Sciences 184:179–197. Moore, P.A., and J.L. Grills. 1999. Chemical orientation to food by the crayfish Orconectes rusticus: influence of hydrodynamics. Animal Behaviour 58:953–963. Naylor, E., and J.A. Atkinson. 1972. Pressure and the rhythmic behaviour of inshore animals. Symposia of the Society for Experimental Biology 26:395–415. Nilsson, D.-E. 1988. A new type of imaging optics in compound eyes. Nature 332:76–78. O’Callaghan, F.E., and P.J. Fraser. 2010. Pressure and temperature related non-linearities in the sensing of angular acceleration in shore crabs. Abstracts of the IXth International Congress of Neuroethology Poster 320. Pabst, H., and D. Kennedy. 1967. Cutaneous mechanoreceptors influencing motor output in the crayfish abdomen. Zeitschrift für vergleichende Physiologie 57:190–208. Page, T.L., and J.L. Larimer. 1976. Extraretinal photoreception in entrainment of crustacean circadian rhythms. Photophysiology 23:245–251. Patek, S.N. 2001. Spiny lobsters stick and slip to make sound. Nature 411:153–154. Patek, S.N. 2002. Squeaking with a sliding joint: mechanics and motor control of sound production in palinurid lobsters. Journal of Experimental Biology 205:2375–2385. Patek, S.N., and R.L. Caldwell. 2006. The stomatopod rumble: sound production in Hemisquilla californiensis. Marine and Freshwater Behaviour and Physiology 39:99–111. Patek, S.N., L.E. Shipp, and E.R. Staaterman. 2009. The acoustics and acoustic behavior of the California spiny lobster (Panulirus interruptus). Journal of the Acoustical Society of America 125:3434–3443. Patullo, B.W., and D.L. Macmillan. 2007. Crayfish respond to electrical fields. Current Biology 17:R83–R84.

81

82

DeForest Mellon, Jr. Patullo, B.W., and D.L. Macmillan. 2010. Making sense of electrical sense in crayfish. Journal of Experimental Biology 213:651–657. Paul, D.H. 1972. Decremental conduction over “giant” afferent processes in an arthropod. Science 176:680–682. Plummer, M.R., J. Tautz, and J.J. Wine. 1986. Frequency coding of waterborne vibrations by abdominal mechanosensory interneurons in the crayfish Procambarus clarkii. Journal of Comparative Physiology A 158:751–764. Prosser, C.L. 1934. Action potentials in the nervous system of the crayfish. II. Responses to illumination of the eye and caudal ganglion. Journal of Cellular and Comparative Physiology 4:363–377. Ripley, S.H., B.M.H. Bush, and A. Roberts. 1968. Crab muscle receptor which responds without impulses. Nature 218:1170–1171. Sandeman, D.C. 1985. Crayfish antennae as tactile organs: their mobility and the responses of their proprioceptors. Journal of Comparative Physiology A 157:363–373. Sandeman, D.C. 1989. Physical properties, sensory receptors and tactile reflexes of the antenna of the Australian freshwater crayfish Cherax destructor. Journal of Experimental Biology 141:197–217. Sandeman, D.C., and J.L. Denburg. 1976. The central projections of chemoreceptor axons in the crayfish revealed by axoplasmic transport. Brain Research 115:492–496. Sandeman, D.C., and A. Okajima. 1972. Statocyst-induced eye movements in the crab Scylla serrata. I. The sensory input from the statocyst. Journal of Experimental Biology 57:187–204. Sandeman, D.C., and R.E. Sandeman. 1996. Pre- and postembryonic development, growth and turnover of olfactory receptor neurons in crayfish antennules. Journal of Experimental Biology 199:2409–2418. Sandeman, D.C., and D. Varju. 1988. A behavioural study of tactile localization in the crayfish Cherax destructor. Journal of Comparative Physiology A 163:525–536. Sandeman, D.C., R.E. Sandeman, and H.G. de Couet. 1990. Extraretinal photoreceptors in the brain of the crayfish Cherax destructor. Journal of Neurobiology 21:619–629. Sandeman, D.C., R.E. Sandeman, C. Derby, and M. Schmidt. 1992. Morphology of the brain of crayfish, crabs and spiny lobsters: a common nomenclature for homologous structures. Biological Bulletin 183:304–326. Sandeman, D.C., B. Beltz, and R. Sandeman.1995. Crayfish brain interneurons that converge with serotonin giant cells in accessory lobe glomeruli. Journal of Comparative Neurology 352:263–279. Schmidt, M. 2007. The olfactory pathway of decapod crustaceans—an invertebrate model for lifelong neurogenesis. Chemical Senses 32:365–384. Schmidt, M., and B.W. Ache. 1996. Processing of antennular input in the brain of the spiny lobster, Panulirus argus. II. The olfactory pathway. Journal of Comparative Physiology A 178:605–628. Schmidt, M., and C.D. Derby. 2005. Non-olfactory chemoreceptors in asymmetric setae activate antennular grooming behavior in the Caribbean spiny lobster Panulirus argus. Journal of Experimental Biology 208:233–248. Schmidt, M., and W. Gnatzy. 1984. Are the funnel-canal organs the “campaniform sensilla” of the shore crab, Carcinus maenas (Decapoda, Crustacea)? II. Ultrastructure. Cell and Tissue Research 237:81–93. Schmidt, M., and DeF. Mellon, Jr. 2011. Neuronal processing of chemical information in crustaceans. Pages 123–147 in T. Breithaupt and M. Thiel, editors. Chemical communication in crustaceans. Springer, New York. Schmidt, M., L. Van Ekeris, and B.W. Ache. 1992. Antennular projections to the midbrain of the spiny lobster. I. Sensory innervation of the lateral and medial antennular neuropils. Journal of Comparative Neurology 318:277–290. Schmiedel-Jakob, I., P.A.V. Anderson, and B.W. Ache. 1989. Whole cell recordings from lobster olfactory receptor cells: responses to current and odor stimulation. Journal of Neurophysiology 61:994–1000. Schöne, H. 1954. Statocystenfunktion und statische Lageorientierung bei dekapoden Krebsen. Zeitschrift für vergleichende Physiologie 36:241–260.



Sensory Systems of Crustaceans

Shaw, S.R., and S. Stowe. 1982. Photoreception. Pages 291–367 in H.L. Atwood and D.C. Sandeman, editors. The biology of Crustacea, Vol. 3, Neurobiology: structure and function. Academic Press, New York. Solov’yov, I.A., D. Chandler, and K. Schulten. 2010. Cryptochrome and magnetic sensing. http://www. ks.uiuc.edu/Research/cryptochrome. Stensmyr, M., S. Erland, E. Hallberg, R. Wallén, P. Greenaway, and B. Hansson. 2005. Insect-like olfactory adaptations in the terrestrial giant robber crab. Current Biology 15:116–121. Steullet, P., D.H. Edwards, and C.D. Derby. 2007. An electric sense in crayfish? Biological Bulletin 213:16–20. Sullivan, J.M., and B.S. Beltz. 2001. Neural pathways connecting the deutocerebrum and lateral protocerebrum in the brains of decapod crustaceans. Journal of Comparative Neurology 441:9–22. Sullivan, J.M., and B.S. Beltz. 2004. Evolutionary changes in the olfactory projection neuron pathways of eumalacostracan crustaceans. Journal of Comparative Neurology 470:25–38. Sullivan, J.M., and B.S. Beltz. 2005. Integration and segregation of inputs to higher-order neuropils in the crayfish brain. Journal of Comparative Neurology 481:118–126. Sullivan, J.M., J.L. Benton, D.C. Sandeman, and B.S. Beltz. 2007. Adult neurogenesis: a common strategy across diverse species. Journal of Comparative Neurology 500:574–584. Sullivan, J.M., M.C. Genco, E.D. Marlow, J.L. Benton, B.S. Beltz, and D.C. Sandeman. 2009. Brain photoreceptor pathways contributing to circadian rhythmicity in crayfish. Chronobiology International 26:1136–1168. Sztarker, J., and D. Tomsic. 2004. Binocular visual integration in the crustacean nervous system. Journal of Comparative Physiology A 190:951–962. Sztarker, J., N.J. Strausfeld, and D. Tomsic. 2005. Organization of optic lobes that support motion detection in a semiterrestrial crab. Journal of Comparative Neurology 493:396–411. Tautz, J., and D.C. Sandeman. 1980. The detection of waterborne vibration by sensory hairs on the chelae of the crayfish. Journal of Experimental Biology 88:351–356. Taylor, R.C. 1975. Physical and physiological properties of the crayfish antennal flagellum. Journal of Neurobiology 6:501–519. Tazaki, K. 1977. Nervous responses from mechanosensory hairs on the antennal flagellum in the lobster Homarus gammarus (L.). Marine Behaviour and Physiology 5:1–18. Tierney, A.J., C.S. Thompson, and D.W. Dunham. 1986. Fine structure of aesthetasc chemoreceptors in the crayfish Orconectes propinquus. Canadian Journal of Zoology 64:392–399. Vaissiere, R. 1961. Morphologie et histologie comparées des yeux des crustacés copépods. Archives de Zoologie Expérimentale et Générale 100:1–125. Vedel, J.P. 1980. The antennal motor system of the rock lobster: competitive occurrence of resistance and assistance reflex patterns originating from the same proprioceptor. Journal of Experimental Biology 87:1–22. Versluis, M., B. Schmitz, A. von der Heydt, and D. Lohse. 2000. How snapping shrimp snap: through cavitating bubbles. Science 289:2114–2117. Vogt, K. 1977. Ray path and reflection mechanisms in crayfish eyes. Zeitschrift für Naturforschung C: Bioscience 32C:466–468. Wachowiak, M., and B.W. Ache. 1994. Morphology and physiology of multiglomerular olfactory projection neurons in the spiny lobster. Journal of Comparative Physiology A 175:35–48. Wachowiak, M., C.E. Diebel, and B.W. Ache. 1996. Functional organization of olfactory processing in the accessory lobe of the spiny lobster. Journal of Comparative Physiology A 178:211–226. Waterman, T.H. 1961. Light sensitivity and vision. Pages 1–64 in T.H. Waterman, editor. The physiology of Crustacea, Vol. 2, Sense organs, integration and behavior. Academic Press, New York. Weatherby, T.M., K.K. Wong, and P.H. Lenz. 1994. Fine structure of the distal sensory setae on the first antennae of Pleuromamma xiphias Giesbrecht (Copepoda). Journal of Crustacean Biology 14:670–685. Weissburg, M.J., and R.K. Zimmer-Faust. 1994. Odor plumes and how blue crabs use them in finding prey. Journal of Experimental Biology 197:349–375.

83

84

DeForest Mellon, Jr. Welsh, J.H. 1934. The caudal photoreceptor and responses of the crayfish to light. Journal of Cellular and Comparative Physiology 4:379–388. Wiersma, C.A.G., E. Furshpan, and E. Florey. 1953. Physiological and pharmacological observations on muscle receptor organs of the crayfish Cambarus clarkii (Girard). Journal of Experimental Biology 30:136–150. Wiese, K. 1976. Mechanoreceptors for near-field water displacements in crayfish. Journal of Neurophysiology 39:816–833. Wilkens, L.A. 1988. The crayfish caudal photoreceptor: advances and questions after the first half century. Comparative Biochemistry and Physiology 91C:61–68. Wine, J.J., and F.B. Krasne. 1982. The cellular organization of crayfish escape behavior. Pages 242–292 in D.C. Sandeman and H.L. Atwood, editors. The biology of Crustacea, Vol. 4, Neural integration and behavior. Academic Press, New York. Yen, J, P.H. Lenz, D.V. Gassie, and D.K. Hartline. 1992. Mechanoreception in marine copepods: electrophysiological studies on the first antennae. Journal of Plankton Research 14:495–512. Young, D. 1989. Nerve cells and animals behaviour. Cambridge University Press, Cambridge. Zucker, R.L., D. Kennedy, and A.I. Selverston. 1971. Neuronal circuit mediating escape responses in crayfish. Science 173:645–650.

4 PERIPHERAL COMPONENTS OF CRUSTACEAN MOTOR SYSTEMS

Harold Atwood Abstract The major peripheral components of crustacean motor systems include excitatory and inhibitory motor neurons, their neuromuscular junctions, and their target muscle fibers. These were extensively investigated during the past century because they are readily accessible for experimental procedures and are relatively large compared with counterparts in other major animal groups. In decapod crustaceans, muscle innervation is remarkable in its economy: a small number of motor neurons supplying each muscle governs a large movement repertoire. This sparse provision of efferent connections is compensated by elaborate diversity among constituent neuromuscular junctions and their partner muscle fibers. Neuromuscular junctions, each comprising many individual synapses, evoke postsynaptic potentials ranging from small-amplitude potentials that often facilitate strongly with repetitive activation, to largeamplitude potentials that typically depress with repetition. Motor neurons specialized for maintained activity usually possess a preponderance of the former type of junction; motor neurons devoted to periodic brief episodes of intense activity are often exclusively endowed with the latter type; while many motor neurons (including those innervating limb muscles) that must operate over a wide range of frequencies to produce highly variable movements supply the constituent muscle fibers of target muscles with junctions that vary greatly in physiological response properties. In some muscles, modulation of both strength and speed of muscular contraction is achieved in part through the influence of peripheral inhibitory neurons, which operate on the postsynaptic muscle membrane to regulate its transmembrane potential, and on excitatory motor terminals to influence the emission of the excitatory transmitter substance. Neuromuscular recruitment of muscle fiber contractions appropriate for the required movement underlies the observed range of locomotory activity. Crustacean muscle fibers are characterized by great variation in contractile properties, arising from differences in membrane excitability, structure, and biochemical endowments; adaptive combinations of these features

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Harold Atwood equip them for a range of physiological roles. Typically, muscle fibers underlying rapid movements have short sarcomeres, high membrane excitability conducive to production of action potentials, and biochemical properties enabling rapid contraction. In contrast, muscle fibers utilized for slow, sustained movements or maintenance of posture have long sarcomeres that permit strong but slow contraction, lower membrane excitability, and biochemical features designed for slow contraction and continuous energy supply. Matching of neuromuscular and muscle fiber properties accounts for the wide range of performance in crustacean muscles. An additional level of modulation is imposed by circulating neurohormones, which affect both neuromuscular junctions and muscle fibers.

INTRODUCTION Large decapod crustaceans have long attracted the attention of physiologists intent on revealing basic mechanisms of nerve conduction, synaptic transmission, and muscular contraction. The large size of their neuromuscular components and their readily identifiable peripheral motor axons stimulated early experimentation. Indeed, starting in the 19th century, the discovery and characterization of individual motor neurons that became known for the characteristic contractions they evoke in their target muscles contributed strongly to the concept of identified neurons in arthropods and other invertebrates (Wiersma 1961). We owe landmark discoveries, including those relating to identification of excitatory and inhibitory transmitter substances, membrane calcium channels, presynaptic inhibition, and various forms of synaptic facilitation and depression, to successful exploration of crustacean neuromuscular systems. Introduction of particularly favorable crustacean preparations such as those from giant muscle fibers of barnacles has stimulated work on membrane ion channels. Comparative physiologists have also found crustacean muscles attractive because of their striking adaptation to a wide variety of habitats and specialized actions. Basic features of crustacean neuromuscular systems will be reviewed with the aim of matching these to the demands of motor performance and, ultimately, of survival. Most is known about neuromuscular mechanisms in the large walking legs and claws of several decapod groups, including especially crayfish and lobsters (Astacidae), and true crabs (Brachyura; Govind and Atwood 1982). Other appendages, including pleopods and uropods, antennae and antennules, eyestalks, and scaphognathites, have also been investigated, though less comprehensively. In the mid-1960s and thereafter, studies of abdominal muscles (axial muscles) of crayfish, clawed lobsters, and spiny lobsters have contributed important insights into neuromuscular features involved in differential movement and postural control, and these muscles have been favorite preparations for teaching purposes. The foregut muscles that operate the gastric and pyloric portions of the stomach of decapod crustaceans have been less studied for neuromuscular physiology than for the nerve impulse patterns that reach them via motor neurons located in the stomatogastric ganglion. In fact, the crustacean stomatogastric system is one of the most heavily investigated small neural systems. In the present account, it will not be considered, as it receives ample attention elsewhere (­chapter 13 in this volume). Here, the focus will be on the neuromuscular systems of limb and axial muscles. The most striking general feature of crustacean (and insect) neuromuscular systems is that they accomplish highly complex movements with muscles innervated by small numbers of motor neurons. This differs from the situation in vertebrates, where much larger numbers of motor neurons are employed to control each muscle. Motor performance in mandibulate arthropods is not inferior to that of vertebrates. In crustaceans, the range and complexity of motor performance is brought about in large measure by adaptive elaboration of the features of constituent muscle fibers in combination with great diversification of neuromuscular synaptic properties, both excitatory and



Peripheral Components of Crustacean Motor Systems

inhibitory. Matching of peripheral synapses with client muscle fibers leads to a wide range of speed and strength among muscles and often among constituent muscle fibers of individual muscles. This major theme dominates the general overview of crustacean neuromuscular systems. More detailed consideration of central control mechanisms for recruitment of neuromuscular activity are considered elsewhere in this volume. Past reviews of crustacean neuromuscular systems, emphasizing different components and processes, provide a rich background for the present account. Comparative neuromuscular mechanisms have been extensively reviewed by Hoyle (1983). Reviews of muscles, innervation, and motor systems are included in volumes 3 and 4 of The Biology of Crustacea (Bliss 1982). Synaptic properties and neuromuscular modulation have been reviewed several times (Atwood 1976, Atwood and Wojtowicz 1986, Atwood and Tse 1993), most recently by Atwood and Klose (2009a, 2009b). Aspects of muscle properties, neuromodulation, neurotransmitter receptor properties, neuromuscular development, and adaptation appear in three volumes on crustacean nervous systems edited by Wiese et al. (1990) and Wiese (2002a, 2002b). Other relevant reviews will be noted in connection with specific topics. Throughout the following survey, functionally relevant features of neuromuscular components will be presented, but not detailed analyses of cellular and molecular mechanisms. The latter are reviewed by Lnenicka (­chapter 6 in this volume).

MOTOR NEURONS Excitatory Neurons Muscle Innervation All crustacean motor neurons, excitatory and inhibitory, provide multiterminal innervation to the muscle fibers they innervate: motor neurons form numerous neuromuscular junctions along the length of each innervated muscle fiber. This contrasts with the predominant pattern of vertebrate skeletal muscles, in which each muscle fiber usually receives only one or two neuromuscular junctions, typically large end-plates. Multiterminal innervation removes the necessity of propagated action potentials in crustacean muscle fibers, although large action potentials often occur. Many crustacean muscle fibers are designed to contract in the absence of action potentials. This range in membrane excitability points to one major factor controlling speed of tension development and total tension in individual muscle fibers. Features of Excitatory Motor Neurons Motor neurons typically have centrally located cell bodies, usually electrically inexcitable and devoid of dendrites. Neuritic processes functioning as dendrites emanate from the major axon as it travels outward to the periphery; these processes receive inputs from sensory neurons and interneurons, and are functionally analogous in many ways to dendrites of vertebrate motor neurons. Marked differences in size and excitation occur among individual motor neurons. Regarding excitation, some motor neurons are tonically active or frequently active, while others are silent most of the time and can be recruited to fire action potentials only in response to intense sensory or interneuronal input. This difference in activity pattern was explored by Kennedy and Takeda (1965a, 1965b) in their landmark investigation of crayfish abdominal muscles, and they designated such neurons as “tonic” and “phasic,” respectively. Tonic neurons to small abdominal muscles regulate posture, while phasic neurons, innervating the large, fast flexor and extensor

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Harold Atwood muscles involved in the escape response, come into action only occasionally, when rapid and powerful activity is called for. Structurally, the tonic abdominal motor neurons and their axons are much smaller than their phasic counterparts, a feature that may contribute to their recruitment for motor activity. The phasic-tonic distinction and terminology has gradually been transported to limb muscles as well. Here, classical work distinguished paired “fast” and “slow” motor neurons supplying several major limb muscles, such as the closer muscle of the crayfish claw; this distinction was based on the type of contraction evoked by the respective neurons (Wiersma 1961). This terminology is still retained, but recognition of the differences in firing patterns, and the effects of these on neuronal properties, has led to frequent adoption of Kennedy and Takeda’s (1965a) terminology. It must be recognized, however, that firing patterns and colligative properties of crustacean motor neurons are very diverse, and that many neurons are intermediate in their properties. For example, the main flexor muscle of the carpopodite in the American lobster Homarus americanus is innervated by four excitatory motor neurons: specialized “fast” (phasic) and “slow” (tonic) motor neurons, and two neurons with intermediate properties (Wiens et al. 1991). Thus, the highly differentiated “phasic” and “tonic” phenotypes are readily distinguished, but they represent well-defined extremes of a continuum. Is size of motor neuron closely linked to excitation properties, as in the “size principle” recognized for mammalian motor neurons? The observations on the crustacean abdominal motor neurons certainly suggest such a relationship (small motor neurons being more readily recruited by excitatory inputs), as do studies on several other systems including lobster pleopod muscles (Hoyle 1983). But for the paired neurons of crayfish claw and leg muscles with dual excitatory innervation, the size principle does not apply: there is no difference in size between the phasic and tonic neurons to the crayfish claw closer muscle, and the tonic excitatory neuron of the walking leg’s main extensor muscle is larger than its phasic partner (Bradacs et al. 1997). Therefore, while neuronal size may contribute to recruitment in some motor neuron pools, it is not a major factor in others. Membrane excitability attributed to different mixes of ion channels imposes a more fundamental differentiation and often plays the major role. Neurotransmitters The primary neurotransmitter substance responsible for the excitatory junctional potentials (EJPs) in the vast majority of crustacean muscles is glutamic acid, while a closely related amino acid, gamma-aminobutyric acid (GABA), is the major neurotransmitter at inhibitory junctions. A few muscles in the foregut receive cholinergic innervation. Work on crustacean neuromuscular systems beginning in the 1960s contributed in a major way toward proving that amino acids function as neurotransmitters (­chapter 6 in this volume). Excitatory neurons often have cotransmitters that serve as neuromodulators through actions on second-messenger systems and muscle membrane ion channels; this will be considered below (Neurohormonal Modulation) (see also ­chapters 5 and 6 in this volume). Synaptic Structure Many ultrastructural studies of crustacean neuromuscular junctions have accumulated. They reveal numerous individual structures along each neuromuscular junction that share features with central synapses in vertebrate and other nervous systems and that will be referred to as individual synapses (Fig. 4.1). Thus, a single neuromuscular junction (contact between a motor nerve ending and a muscle fiber) often forms scores of individual separated synapses. Typically, they occur on varicosities of the nerve terminal. Each synapse is relatively small (typically about 1 µm 2 in contact area) and consists of closely apposed, densely staining pre- and postsynaptic



Peripheral Components of Crustacean Motor Systems

Fig. 4.1. Motor innervation of crustacean muscle fibers. (A)  A  terminal varicosity with multiple individual excitatory synapses (crayfish opener muscle). The excitatory nerve terminal, plentifully supplied with mitochondria and synaptic vesicles, forms five clearly discernible synapses (S), of which three possess presynaptic dense bodies, which are components of the “active zones” of these synapses (asterisks). Scale bar: 0.5 µm. (B) Inhibitory and excitatory synapses in stretcher muscle of the crab Hyas areneus. An inhibitory nerve terminal (In) forms an axoaxonal synapse (a) on an excitatory nerve terminal (Ex), which in turn forms a neuromuscular synapse (nm) on the muscle fiber (M). The inhibitory terminal also forms a neuromuscular synapse on the muscle fiber. All three of the designated synapses possess presynaptic dense bodies (active zone components), around which synaptic vesicles cluster. Scale bar: 0.5 µm (adapted from Atwood 1982, with permission from Academic Press).

membranes. Individual synapses may also have one or more electron-dense presynaptic “dense bodies” acting as focal points for clustered synaptic vesicles at putative release sites. Some individual synapses lack a dense body, and these also lack a cluster of synaptic vesicles. Such observations indicate that individual synapses vary in their ability to release neurotransmitter on activation; those lacking a dense body are likely “silent” (Atwood and Wojtowicz 1999). While the probability of transmitter release by individual synapses may be low, there is very good evidence that more synapses can be recruited for neurotransmitter release during facilitation of transmission by bursts of impulses or sustained stimulation, in both crustaceans and insects (Wojtowicz et al. 1994, Quigley et al. 1999, Peled and Isacoff 2011). Individual crustacean synapses studied with freeze-fracture electron microscopy display limited numbers of characteristic large presynaptic membrane particles, which are thought to be Ca 2+ channels responsible for fast fusion and exocytosis of synaptic vesicles. Their number and distribution does not differ significantly between phasic and tonic neurons investigated to date (Msghina et al. 1999). However, pharmacological evidence for different types of presynaptic Ca 2+ channels in phasic and tonic motor axons of a crab leg muscle has appeared (Rathmayer

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Harold Atwood et  al. 2002a). While both phasic and tonic terminals express P/Q-type Ca 2+ channels, additional N-type channels occur only on phasic terminals, and R-type channels on tonic terminals. However, this difference in Ca 2+ channel mix cannot explain the large difference in transmitter release per synapse in the two terminals (see review by Atwood and Karunanithi 2002). Interestingly, the general structure of individual synapses on phasic and tonic motor neurons does not differ; hence, the marked differences in neurotransmitter output at phasic and tonic neuromuscular junctions is not due to definable ultrastructural differences. However, the nerve endings of the neuromuscular junction do show gross phasic-tonic structural differences: those of phasic excitatory neurons are thin and filiform, while those of tonic neurons are larger and varicose, with larger populations of synaptic vesicles (Atwood and Karunanithi 2002, ­chapter 6 in this volume). The prominent varicosities of tonic neurons are well endowed with mitochondria, and this correlates well with their much greater resistance to synaptic depression (Lnenicka and Morley 2002). Synaptic Variation Intracellular records of EJPs can be characterized according to their initial amplitude and the changes that occur with repetitive stimulation. Initial amplitude is governed by the electrical properties of the muscle fiber membrane and by the amount of neurotransmitter released by a single motor nerve action potential at responsive synapses (Cooper et al. 1996), while changes in amplitude with repeated stimulation reflect presynaptic changes leading to more transmitter release (facilitation) or to decline in transmitter release (synaptic depression). The mechanisms of transmitter release and facilitation are reviewed by Lnenicka (­chapter 6 in this volume). Excitatory junctional potentials vary enormously from one muscle to another and also within an individual muscle (Figs. 4.2 and 4.3). The fast abdominal muscles of crayfish and lobster exhibit large (10 mV or greater) EJPs for single impulses, which often trigger muscle action potentials of variable amplitude and shape, leading to a fast twitch contraction of the muscle. In contrast, the tonic motor axons of the small abdominal postural muscles produce small EJPs (usually less than 10 mV), which vary in amplitude among the constituent muscle fibers. In limb muscles, complexity is the rule. Synaptic diversity within muscles and among species was initially reported by Hoyle and Wiersma (1958), and subsequent studies have confirmed and elaborated this general picture. As an example, the fiber-by-fiber analysis of the leg closer muscle of a crab (Eriphia spinifrons) by Rathmayer and Erxleben (1983) shows large variation in EJP amplitude and shape for both excitatory motor axons innervating this muscle (Figs. 4.2A and 4.3A). The synaptic responses are matched to muscle fiber properties: fibers that receive innervation mainly from the fast (phasic) closer excitor exhibit rapid decay of EJPs due to short time-constants of the muscle fiber membrane, while fibers heavily innervated by the slow (tonic) excitor decay much more slowly due to longer muscle membrane time-constants. Other muscle fiber properties also vary, providing a basis for classification of four fiber types (Fig. 4.2A and Table 4.1). Biochemical features will be outlined in a later section. This highly refined variation in synaptic properties within a single muscle (typical of many crustacean limb muscles) accounts in part for the range and precision of muscle performance: specific sets of muscle fibers can be recruited by peripheral synapses in accordance with changes in nerve activity patterns. Differential recruitment of muscle fibers is evident also in limb muscles that receive only a single excitatory motor neuron; the “opener” muscle of the crayfish claw and walking leg is the most thoroughly studied example. In decapods, the “opener” muscle shares its single motor axon with the functionally distinct “stretcher” muscle in the carpopodite joint, and independent use of these muscles is made possible by their separate inhibitory innervation (Fig. 4.4). Synaptic potentials evoked in individual muscle fibers by the opener-stretcher motor axon differ greatly

A

P P T

T

P T

T FCE

B

7

8

6

9

CI

SCE

5 4 3 2 1

200 µm i

ii

iii

iv

Fiber group

Fig. 4.2. Innervation and muscle fiber types in crustacean limb muscles. (A)  Diagrammatic representation of excitatory motor innervation in a muscle with one tonic-type (T) axon (left), and in a muscle with two excitatory motor axons, one tonic (T) and one phasic (P) axon (right). In both cases, the axons innervate a range of muscle fiber types, indicated by differences in sarcomere length. Muscle fibers with short sarcomeres contract more rapidly than those with long sarcomeres. Characteristic differences in the EJPs are illustrated. In a doubly innervated muscle (ii), large EJPs generated by the phasic axon may trigger graded or all-or-nothing action potentials. Some fibers are innervated exclusively by one of the two axons, but the majority are innervated by both (from Atwood 1976, reproduced with permission from Elsevier). (B) Diagram of muscle fiber types and innervation in a crab closer muscle. Innervation by a phasic excitatory axon (FCE, “fast closer excitor”), a tonic excitatory axon (SCE, “slow closer excitor”), and common inhibitory axon (CI) is shown. Four muscle fiber types are distinguished on the basis of their characteristic electrophysiological and histochemical features (see Table 4.1). Individual identified muscle fibers are numbered (from Rathmayer and Maier 1987, reproduced with permission from Oxford University Press).

Type II medium medium small, facilitating medium, some facilitation weak high high fast oxidative glycolytic intermediate

medium, facilitating medium, no facilitation strong low medium slow oxidative slow

Inhibitory innervation Myosin ATPase activity Mitochondrial enzymes Metabolic type

Contraction type

low long

Type I

Parameter Membrane potential Membrane time constant EJPs, tonic axon EJPs, phasic axon

fast oxidative glycolytic fast

high

high

none medium, facilitating none

Type III high short

fast

fast glycolytic

low

high

none small, facilitating none

Type IV high short

Table 4.1.  Muscle fiber types in leg muscles of the crab Eriphia spinifrons, based on combinations of electrophysiological, biochemical, and contractile features (after Rathmayer and Maier 1987, with permission from Oxford University Press)



Peripheral Components of Crustacean Motor Systems

in amplitude and facilitation properties, resembling in most respects those of the tonic motor axons of the antagonistic “closer” and “bender” muscles. Fast and slow contractions are readily apparent in the muscles with a single motor axon (Wiersma 1951), due to selective recruitment of rapidly contracting and slowly contracting muscle fibers, respectively, by different impulse patterns (Fig. 4.4B; Atwood 1973, 1976). Inhibitory motor neurons aid in selective recruitment of different muscle fiber types, as will be discussed in a later section. Repetitive stimulation generally shows that neuromuscular transmission facilitates initially for both tonic and phasic excitors (Figs. 4.2B and 4.3B). However, when stimulation is continued, depression soon sets in at phasic, but not at tonic, neuromuscular junctions (Fig. 4.3B; ­chapter 6 in this volume). In some phasic motor neurons, two types of depression appear: short-term or low-frequency depression, evident after the first stimulus and progressing with subsequent stimuli delivered at low frequency; and high-frequency depression, usually evident at stimulation frequencies of 5 Hz or more. Depletion of neurotransmitter stores and slow replenishment linked to limited energy metabolism can account for high-frequency depression (Nguyen et al. 1997). Low-frequency depression is attributed to action of neural phosphatases (Silverman-Gavrilla et al. 2005). The functional implication of this propensity for depression is that most phasic excitors cannot reliably maintain optimal transmission over long periods of activity (Fig. 4.3B). They are designed for high initial performance, as required during escape, extreme exertion, or bursts of fast locomotion. Tonic excitors, with initially modest transmitter release, resist depression and often show marked facilitation with repeated stimulation. Even in leg muscle fibers innervated by both phasic and tonic axons, EJPs of the latter invariably display more pronounced facilitation (Fig. 4.3A). Tonic excitors do most of the work during normal locomotion, as revealed by implanted electrodes in freely moving animals. They are endowed with more mitochondria in synaptic varicosities, indicative of a better energy supply for maintaining neurotransmission. In addition, their terminals release less transmitter than those of a phasic excitor when both are exposed to a defined level of intracellular calcium, demonstrating lower calcium sensitivity of the transmitter-releasing apparatus (Millar et al. 2005). A general model to account for phasic-tonic differences in neurotransmission has recently been published (Pan and Zucker 2009). Inhibitory Neurons Muscle Innervation by Inhibitory Motor Neurons Most limb, appendage, and axial muscles are innervated by a single inhibitory motor axon, but an important exception is found in the distal “opener” and “stretcher” muscles of walking legs and claws, which receive two inhibitory axons (Fig. 4.4). The distribution of inhibitory innervation within target muscles is usually widespread, affecting most muscle fibers to varying degrees; but some limb muscle fibers that contract rapidly appear to receive little or no inhibitory innervation. The two inhibitory neurons innervating the opener-stretcher “motor unit” are termed “specific” and “common” inhibitors, on the basis of their distributions: specific inhibitors innervate only the opener or the stretcher muscle, while the common inhibitor innervates all of the muscles operating a limb (Fig. 4.4B). Thus, the specific inhibitors allow the opener and stretcher muscles to be used separately, while the common inhibitor affects all limb muscles and serves to modify their contraction speed (Wiens 1989). Postsynaptic and Presynaptic Inhibition Muscles with inhibitory innervation exhibit inhibitory postsynaptic potentials (inhibitory junctional potentials:  IJPs) with intracellular recording (Fig. 4.4B). Inhibitory potentials may be either hyperpolarizing or depolarizing, depending on the relationship between the muscle fiber’s membrane potential and the reversal potential for inhibitory transmitter action (which usually is close to the resting membrane potential of the muscle fiber). Some fibers with weak inhibitory

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B: Phasic Axon

A Fiber 1

Type IV

2 Type I

3 4

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Type II

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7 8 9

Type IV

B 5 Hz

Phasic

0

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0

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2 mV

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2 msec 4

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Fig. 4.3. Variation in EJPs of phasic and tonic motor axons. (A) EJPs recorded from identified muscle fibers in the closer muscle of a crab (Eriphia spinifrons). The numbered muscle fibers and their fiber types are shown schematically in Figure 4.2B. Responses to stimulation of the “tonic” excitatory neuron appear in column A (Tonic), and corresponding responses to the “phasic” neuron evoked by the same pattern of stimulation appear in column B (Phasic). Note that Type III and Type IV fibers show minimal or no response to the tonic neuron, while all muscle fibers are innervated by the phasic neuron. Facilitation during repetitive stimulation is always greater for tonic than for phasic EJPs in a given muscle fiber. Calibration pulses at the start of each trace are 2 mV and 20 msec (from Rathmayer and Hammelsbeck 1985, adapted with permission from The Company of Biologists). (B) Facilitation and fatigue in a tonic (bottom) and a phasic (top) neuron in abdominal muscles of a crayfish (Procambarus clarkii). EJPs of the tonic neuron stimulated continuously for 20 min facilitated throughout the stimulation period, while EJPs of the phasic neuron, occurring at half the frequency over the same time span, exhibited progressive synaptic depression (from Nguyen et al. 1997, with permission from The American Physiological Society).

input, or in which the inhibitory reversal potential matches the resting membrane potential, may not show IJPs unless the membrane potential is displaced from its resting value. The main effect of inhibitory transmitter action is to increase postsynaptic membrane conductance and to direct the membrane potential toward the reversal potential for inhibitory transmitter receptor



Peripheral Components of Crustacean Motor Systems

channels, which are predominantly permeable to the chloride ion. These receptors are ionotropic receptors (GABA A receptors) for the inhibitory neurotransmitter GABA. The increase in membrane conductance invariably reduces the membrane time constant of the affected muscle fiber, shortening EJPs, making their summation less effective and often reducing their amplitudes, thereby weakening or eliminating the muscle fiber’s contraction. In muscles fibers that A

OI OE=SE

SI F S

O

RE

R

CI AF

C B

E B

Flexor

Acc. Flexor Bender Opener

o.n

c.n. Extensor Stretcher

Closer

Fig. 4.4. Innervation of decapod crustacean limb muscles. (A)  Innervation of the distal muscles of the lobster (Homarus americanus) leg, showing the specific inhibitory neurons (OI and SI) of the opener (O)  and stretcher (S)  muscles, respectively, and the common inhibitory neuron (CI) supplying all of the distal muscles. Excitatory neurons are indicated for the opener and stretcher muscles (sharing a single excitatory neuron, OE = SE); the closer (C), bender (B), and extensor (E) muscles (each innervated by two excitatory neurons, one phasic, and one tonic); the main flexor (F) muscle (innervated by four excitatory neurons, one of which, designated RE, is shared with the small rotator (R) muscle); and the accessory flexor (AF) muscle (innervated by a single excitatory motor neuron) (adapted from Wiens 1989 with permission from John Wiley & Sons). (B) Innervation of distal leg muscles in the crab Eriphia spinifrons shown by recording inhibitory junction potentials from all seven muscles in one preparation while stimulating distal branches of the CI neuron in the closer nerve (c.n.) and opener nerve (o.n.); identical results were observed for each nerve branch. Calibration pulse (beginning of each trace), 1 mV, 50 msec. (From Wiens and Rathmayer 1985, reproduced with permission from Springer Science and Business Media).

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Harold Atwood produce graded or all-or-none action potentials (as in fast abdominal muscles), inhibition often eliminates them, leaving the underlying EJP, which may not be sufficient to produce a contraction (review: Atwood 1982). Synapses subserving postsynaptic inhibition are readily identified in electron micrographs, and their structure is generally similar to that of excitatory synapses (Fig. 4.1B). Many inhibitory synapses of limb muscles have larger contact areas and more presynaptic dense bodies (“active zones”) than their excitatory counterparts. This correlates with the larger estimated quantal content of inhibitory neuromuscular transmission, especially for specific inhibitors: for a single nerve impulse, more neurotransmitter is released at an inhibitory neuromuscular junction than at an excitatory one on the same muscle fiber. In some decapod limb muscles, presynaptic inhibition also occurs (Dudel and Kuff ler 1961, ­chapter 6 in this volume). This is manifested by reduction of EJPs when an inhibitory nerve impulse is timed to arrive just before an excitatory one. EJPs decrease due to less neurotransmitter release at the excitatory synapses. Inhibitory synapses responsible for this action (axoaxonal synapses) have been located by electron microscopy on excitatory nerve terminals (Fig. 4.1B), and the physiological effects have been studied with intracellular recording from the nerve terminals and through modeling of inhibitory transmitter action (see review by Atwood and Tse 1993). In brief, the axoaxonal synapses cause an increase in chloride conductance in the excitatory nerve terminal, which decreases the amplitude of the excitatory nerve action potential, especially in distal extremities of the excitatory neuromuscular junction, where many of the excitatory synapses are located. This leads to reduced neurotransmitter output. An additional component of presynaptic inhibition is attributed to participation of G-protein coupled receptors (GABA B receptors) on excitatory nerve terminals (Miwa et al. 1990). About 10–25% of the presynaptic inhibitory action in opener and closer muscles of the crab Eriphia is attributed to GABA B receptors (Rathmayer and Djokaj 2000). Whether activation of these receptors acts on electrical events in the excitatory nerve terminals or more directly on excitation-secretion coupling is still not clear. The GABA B receptors are thought to open channels selective for potassium ions. They have not been found on muscle fibers, and are probably not involved in postsynaptic inhibition. They may be located extrasynaptically on excitatory nerve terminals, and probably prolong the duration of inhibition. Neurotransmitter and Receptors The case for GABA as the major inhibitory neurotransmitter in crustaceans was definitively established in the late 1960s (review:  Atwood 1982). Subsequently, considerable work on the GABA receptors of crustacean muscles and nerve terminals has been undertaken (reviews: Atwood and Tse 1993, Rathmayer and Djokaj 2000). Aspects of GABA action and crustacean GABA receptors are dealt with by Lnenicka (­chapter 6 in this volume). Synaptic Variation Inhibitory neuromuscular junctions exhibit intramuscular variation resembling that of excitatory junctions. In fact, both initial amplitude and short-term synaptic facilitation of EJPs and IJPs in muscles innervated by specific inhibitors are often “matched” in the individual muscle fibers throughout the muscle (Atwood and Bittner 1971). In contrast, IJPs of the common inhibitor are less well matched in their properties to EJPs, and generally exhibit more short-term facilitation (Fig. 4.4B). In muscles with separate phasic and tonic excitatory neurons the common inhibitor is less effective in reducing the contractions evoked by the former, due to smaller inhibitory effects on the most rapidly contracting muscle fibers.



Peripheral Components of Crustacean Motor Systems

Presynaptic inhibition is usually more effective for specific than for common inhibitory motor axons in muscles with dual inhibitory innervation. Nevertheless, the common inhibitor does exert presynaptic inhibition at both phasic and tonic excitatory junctions in a crab closer muscle (Rathmayer and Djokaj 2000). In crayfish and lobster abdominal muscles, presynaptic inhibition has not been found. Even in walking legs, its presence is variable, being most prominent in true crabs (Brachyura), and possibly absent in several leg muscles of the American lobster. Such variation may be attributed to functional requirements in different groups of decapod crustaceans; however, these features have as yet been studied in relatively few species.

MUSCLE FIBERS Structure Sarcomeres Striated muscle fibers are the building blocks of all crustacean muscles, and they are composed of sarcomeres (the repeated structural unit delimited by thin dense Z-lines to which actin filaments are attached). In different muscle fibers, sarcomeres often differ greatly in structure and in functional properties. An easily measured feature is sarcomere length, which is more variable in crustacean muscles than elsewhere in the animal kingdom. In the 1960s and 1970s, studies on various muscles led to the generalization that muscle fibers with short sarcomeres (2–3 µm) are rapidly contracting, and those with long sarcomeres (7–µm) are slowly contracting. On the other hand, the latter can produce much greater maximal tension per unit cross-sectional area (reviews: Atwood 1972, Chapple 1982, Hoyle 1983). This feature is attributed to the longer myofilaments and their greater overlap in longer sarcomeres, in accordance with the sliding-filament model of muscle contraction. Slower contraction speed of long-sarcomere fibers is due in part to fewer sarcomeres in series. While other features of the muscle fiber (especially membrane excitability and biochemical properties) contribute to contraction speed, the relationship between maximal force and sarcomere length has been reaffirmed in a comparative crustacean study, which concluded that muscle or myofilament properties other than sarcomere length need not be invoked to explain the high tensions that can be generated by crustacean claw muscles (Taylor 2000). Membrane Systems Myofibrils within the muscle fiber are separated by a fenestrated internal membrane system, the sarcoplasmic reticulum (SR). In crustacean muscles, junctions between this system and T-tubules that invaginate from the surface typically occur at the interface between actin and myosin filaments (the A-band and I-band interface). At this location, external microelectrode stimulation is effective in producing localized sarcomere contractions, leading to the conclusion that excitation-contraction coupling is mediated by T-tubule junctions with SR here. As in striated muscle fibers of vertebrates, excitation-contraction coupling is thought to involve release of Ca 2+ from terminal cisternae of the SR. Calcium-induced calcium release is likely to be an important mediator of the process (Goblet and Mounier 1986). In addition to its role in excitation-contraction coupling, the SR acts to sequester Ca 2+ rapidly after excitation; this function is essential for relaxation of contraction. In muscle fibers specialized for very rapid contraction and relaxation, the SR’s relative volume is increased. The most extreme known example of enlarged SR in crustaceans occurs in the remotor muscle of the second antenna of the American lobster, a muscle responsible for sound production that

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Harold Atwood can contract at frequencies of 100 Hz or greater (Mendelson 1969). In fast fibers of this muscle, the SR occupies about 70% of the muscle fiber volume, and the contractile myofibrils are consequently greatly reduced in comparison with other muscle fibers (Rosenbluth 1969). The larger relative volume of SR is estimated to increase the rate of Ca 2+ uptake to promote rapid relaxation. An additional invaginating membrane system of tubules, referred to as Z-tubules due to their localization at sarcomere Z-lines, has been described in long-sarcomere fibers, but not in short-sarcomere fibers. Localized stimulation does not produce contraction here, even though small membranous structures interconnect Z- and T-tubules (Franzini-Armstrong et al. 1986). The function of Z-tubules has not been clarified to date, but these tubules do contribute to the electrical properties of the muscle fiber membrane. The extensive membrane systems and additional deep sarcolemmal clefts of crustacean muscle fibers are responsible for their relatively large membrane electrical capacitance. This feature endows many muscle fibers (particularly the slowly contracting ones) with long membrane time constants, and EJPs of such fibers typically have a long time course with slow decay. Biochemical Properties Muscle Fiber Typing With the advent of histochemical methods for muscle fiber typing in mammalian muscles in the 1960s, and adaptation of these procedures for crustacean muscle, it became possible to analyze and classify crustacean muscle fibers according to their biochemical endowments. Numerous studies employing this approach appeared in the 1970s and thereafter. Crustacean muscle fibers were found to vary widely in biochemical features, adding another layer of diversity to that imparted by differences in sarcomere characteristics, and pointing to further functional distinctions. Myofibrillar adenosine triphosphatase (ATPase) activity was widely adopted as a primary criterion for distinguishing fast and slow muscle fibers: fibers with intense staining (high level of myofibrillar ATPase activity) were identified as fast fibers. In conjunction with this, staining for the mitochondrial enzyme NADH diaphorase was adopted to distinguish muscle fibers with high and low mitochondrial content, linked respectively to fatigue resistance and rapid fatigue, respectively. Typical results of this approach are shown for the power-stroke swimming muscles at the base of the modified pereopods in the portunid crab Callinectes sapidus (Fig. 4.5). Color differences are linked to mitochondrial content:  striking differentiation of fast, mitochondrial-poor (white) and slower, mitochondrial-rich (deep pink) muscle fibers, together with intermediates, is readily apparent. Contraction speeds of individual fibers tested with contraction-inducing injection of current through an intracellular microelectrode confirmed the fast-slow functional distinction. Of particular interest in this case, the sarcomere lengths of the two fiber types are relatively short, and not greatly different (around 4 µm in both types), indicating that the difference in contraction speed is due primarily to the difference in ATPase activity (Tse et al. 1983). However, both fiber types in this muscle are faster than the fibers with very long sarcomeres (7–15 µm) found in other crustacean muscles, which invariably also have low myofibrillar ATPase activity. The occurrence of fast-oxidative muscle fibers (with high myofibrillar ATPase and high mitochondrial content: fast and fatigue resistant) has been highlighted in studies of continuously active maxilliped muscles in portunid crabs (Silverman and Charlton 1980). Increasingly refined histochemical and enzymatic analyses at the level of single muscle fibers have been carried out, as exemplified by studies of Rathmayer and colleagues on identified single fibers of the superficial layer in the closer muscle of the crab Eriphia (Rathmayer and



Peripheral Components of Crustacean Motor Systems

Fig. 4.5. Muscle fiber types in the levator muscle of the swimming leg (branches E and F) in a portunid crab (Callinectes sapidus) shown with histochemical stains. Panels I and III show results for NADH diaphorase, (a mitochondrial indicator), while Panels II and IV, taken from an adjacent section of the same muscle, show results for myofibrillar ATPase (an indicator of rate of tension development). Fibers with high mitochondrial density are deep pink in gross coloration (d); fibers with intermediate mitochondrial density are light pink or intermediate (i); and fibers with low mitochondrial density are white (w). Small arrows denote small oval fibers that do not stain. Scale bars: I and II, 0.4 mm; III and IV, 0.1 mm (from Tse et al. 1983, with permission from Canadian Science Publishing).

Maier 1987). With addition of results for other metabolic enzymes, together with observations on innervation, membrane properties, and contraction, four different fiber types were defined (Table 4.1); and even within fiber types, additional variation was apparent. In general, the fast fibers, innervated primarily by the phasic motor axon (Fig. 4.3A), rely on glycolytic metabolism and presumably cannot sustain prolonged contractions; while slow fibers, with prominent innervation by the tonic excitor, are oxidative and presumably fatigue resistant. As Rathmayer and Maier (1987) note: “This makes the muscle a very heterogeneous tissue, in which each fiber might represent a typical, separate metabolic entity.” With advances in molecular analysis, additional biochemical differences have come to light in selected muscles (review: Silverman et al. 1987). Molecular differences in contractile proteins (myosin) and their regulators (troponin, tropomyosin) have been used to distinguish between slow and fast muscle fibers in American lobster walking legs and claws (Mykles 1985, Silverman et al. 1987). The fact that some fibers in these muscles with similar sarcomere lengths and myofibrillar ATPase contract at different rates may be attributed to differences in isoforms of contractile and/or regulatory proteins. This theme has been extended in studies

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Harold Atwood of the slow abdominal f lexor muscles of the Norway lobster, Nephrops norvegicus (Holmes et al. 2002). In this muscle, the fibers are all slow (in comparison to the fast f lexor muscles), but two fiber types can be distinguished: a faster type (S1) that has higher ATPase activity, and a slower type (S2) that has less ATPase activity and expresses an isoform of troponin T (T1 isoform) that is not present in S1 fibers. Tests with isolated, skinned muscle fibers revealed slower kinetics for activation and relaxation, and higher sensitivity to Ca 2+, for the S2 fibers. Thus, isoform differences may account for the different rates of tension development reported within fiber populations with similar sarcomere lengths (Silverman et al. 1987). In addition, the arrangement of these two types of fiber in the slow f lexor muscles provides the basis for tonic tension maintenance (S2 fibers) and slow abdominal movements requiring additional force (S1 fibers). Contraction Membrane Potential Threshold The groundbreaking classical studies of Orkand (1962a, 1962b), in which depolarizing or hyperpolarizing currents were passed into single crayfish muscle fibers while membrane potential and tension were recorded, demonstrated unequivocally that excitation-contraction coupling in crustacean muscle fibers is strictly dependent on the absolute value of the membrane potential, with more tension appearing as it is depolarized past a “threshold” value (Fig. 4.6A). These findings effectively obliterated earlier hypotheses linking excitation-contraction coupling to direct effects of transmitter substances on the contractile machinery (Wiersma 1961). Subsequent studies using versions of Orkand’s technique led to additional findings. First, the value of the threshold membrane potential for excitation-contraction coupling (E-C threshold) has widely different values within populations of muscle fibers, as do fibers’ resting potentials (Atwood et al. 1965, Hoyle 1983). Fibers were discovered in which the E-C threshold is only a few millivolts more positive than the resting potential, so that a very small depolarization is sufficient to elicit tension. In muscles of portunid crabs, cases of fibers with E-C threshold more negative than the resting potential were also disclosed; these fibers were in a state of weak contracture even at rest (Hoyle 1968a, 1968b). In the majority of crustacean muscle fibers, substantial depolarization is needed to reach the E-C threshold, with fast fibers typically requiring more depolarization to contract than slow fibers. Although the mechanisms for the diversity of E-C thresholds are not well understood, available evidence indicates that excitation-contraction coupling in crustacean muscle fibers is totally dependent on external Ca 2+ (unlike vertebrate striated muscle fibers, which can contract for a while even in the absence of external Ca 2+). Depolarization opens Ca 2+ channels and the resulting influx of Ca 2+ can have several effects. First, it can directly induce at least some mechanical activation (seen when other mechanisms are pharmacologically blocked). Second, it can activate the SR rapidly through calcium-induced Ca 2+ release. Third, it may trigger release of Ca 2+ from an inositol 1,4,5-trisphosphate sensitive store, presumably located in the SR (Rojas et al. 1992). Variation in sensitivity of any of these mechanisms could contribute to the E-C threshold. This source of diversity among crustacean muscle fibers remains to be more completely explained. For example, fast fibers, with a more elaborate transverse tubular system, may depend more heavily on entry of external calcium than slow fibers (Ushio and Watabe 1993), but such proposals have yet to be tested with conclusive experiments. In addition to contractions mediated by depolarization, relaxation can be evoked by hyperpolarization of a contracting fiber. Artificially imposed hyperpolarization causes relaxation in fibers that are in a state of resting tension or weak contracture (Hoyle 1968b). As shown by

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Fig. 4.6. Depolarization and tension in crustacean muscle fibers. (A) The membrane potential threshold for contraction, and tension as a function of depolarization, in a crayfish muscle fiber. In each record, upper trace is depolarization caused by injected positive current, and lower trace is the tension response. Resting membrane potential was 75 mV. The fiber must be depolarized by about 20 mV to develop initial tension; above this threshold, tension response increased greatly for small increases in depolarization (from Orkand 1962a, with permission from John Wiley & Sons). (B) Rates of tension development with direct depolarization of single lobster muscle fibers, main leg extensor muscle, illustrating the difference in tension responses between “fast-follower” (i) and “slow-follower” (ii) muscle fibers directly activated by similar depolarizing pulses. In each series, the applied current was progressively increased. Initial peaks in depolarization (upper traces in each record) are due to delayed rectification; tension responses (lower traces in each record) follow these peaks in fast-follower muscle fibers, but not in slow-follower muscle fibers. Tension increases continuously during the depolarization pulse in slow-follower muscle fibers, and often peaks after the pulse has been terminated. Calibrations: time 0.2 sec; voltage 10 mV; tension 20 mg. Resting membrane potentials were: A, 70 mV; B, 65 mV (from Jahromi and Atwood 1971, courtesy of The Wistar Institute, Wistar Archives Collections, Philadelphia, PA).

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Harold Atwood Orkand (1962b) and by Atwood et al. (1965), the inhibitory transmitter substance GABA, and stimulation of the inhibitory motor neuron to effect postsynaptic inhibition, produce relaxation of tension, or prevent it from occurring, solely by regulating the muscle fiber’s membrane potential. Rate of Contraction Muscle fibers subjected to square pulses of depolarizing current to bring their membrane potential past the excitation-contraction coupling threshold exhibit a large range of contraction rates, even when sarcomere length is not markedly different (Fig. 4.6B). Originally, muscle fibers with different contraction speeds were classed as “fast-follower” and “slow-follower” (Atwood et al. 1965). Fibers with very long sarcomeres are mostly “slow-followers,” and fibers in fast abdominal muscles with very short sarcomeres are invariably “fast-followers.” As biochemical information about crustacean muscle fibers increased, it became apparent that some of the differences that had turned up in earlier physiological studies can most reasonably be attributed to differences in biochemical properties of contractile proteins and their regulators, as already related (e.g., Holmes et al. 2002). But additional factors also contribute: effectiveness of Ca 2+ release and uptake by the sarcoplasmic reticulum; and membrane electrical properties and excitability, among others. Membrane Excitability Given the continuous membrane potential-tension relationship (Fig. 4.6A), muscle fibers can produce any degree of tension between threshold and the maximum attainable, depending on the depolarization above E-C threshold imposed by neural or artificial stimulation. Thus, EJPs by themselves can produce graded tension in a muscle fiber, since they can depolarize the fiber to varying extents depending on the impulse frequency in the motor neuron (Fig. 4.7B). Graded summation and facilitation of EJPs depends strongly on incoming impulse patterns. In principle, there is no absolute requirement for additional depolarizing electrogenesis such as muscle action potentials. Many slow muscle fibers are, in fact, electrically inexcitable and do not generate action potentials, even when strongly depolarized. Nevertheless, the capacity to generate muscle action potentials exists in many crustacean muscle fibers, especially fast-acting ones. This is particularly evident in fast abdominal flexor and extensor muscles of crayfish and lobsters (Kennedy and Takeda 1965a) and in some leg muscle fibers (Fig. 4.7A; Hoyle and Wiersma 1958, Atwood et al. 1965). The function of action potentials is to evoke fast contractions for rapid movements. An example in Figure 4.7B illustrates, in a single muscle fiber, slowly developing tension in response to EJPs, with transition to rapidly developing tension when a graded spike is triggered. When action potentials do occur, as in this fiber, they are often graded in amplitude, depending on the rate of rise and amplitude of the depolarization evoking them. However, many fast muscle fibers do react with all-or-none action potentials when depolarization exceeds the threshold membrane potential for triggering an action potential (Fig. 4.7A). Crustacean muscle fibers are dependent on membrane Ca 2+ channels for production of action potentials, as there are no voltage-gated sodium channels available. In fact, much of the initial work on membrane Ca 2+ channels, now known to occur in many cell types, was carried out on crustacean muscle fibers. In general, whether graded or all-or-none electrogenesis is evoked depends on the membrane’s ratio of calcium channels and repolarizing potassium channels of various types. In comparison with action potentials in vertebrate muscle fibers, those of crustacean muscle fibers are slower in time course, more variable, and usually nonpropagated due to distributed innervation (Hoyle 1983). Nevertheless, they have an important function in fast and intermediate muscle fibers, contributing in a major way to the speed and amplitude of the actual contractile event by causing a large influx of external calcium ions.



Peripheral Components of Crustacean Motor Systems

Fig. 4.7. Electrical responses and tension development in single crab muscle fibers responding to (A) direct depolarization and to (B) excitatory EJPs. (A) Electrical and mechanical responses of single muscle fibers. One fiber showed no tension response until an action potential was elicited at a discrete threshold; a second fiber developed variable tension responses when graded membrane responses appeared (adapted from Atwood et al. 1965, with permission from The Physiological Society). (B) Single innervated muscle fibers of a crab (Cancer magister) responding to excitatory nerve stimulation. (i)  EJPs in a fiber of the stretcher muscle produced a small tension response, which abruptly increased when graded action potentials were elicited. Calibration: 1 sec; 20 mV; 0.1 g. (ii) Tension development in a passively responding slow-follower muscle fiber of the closer muscle, with elicitation of EJPs by the tonic axon. Tension appeared with small depolarizations and was graded according to the total depolarization, which increased with frequency of stimulation, and continued to develop after cessation of stimulation. Stimulation rates: 25, 30, 50, 60, 75, 100 Hz. In these records, tension responses are shown as downward deflections in the upper traces. Calibration: 100 msec, 20 mV, 0.2 g (from Atwood et al. 1965, with permission from The Physiological Society).

FUNCTIONAL ORGANIZATION General Principles The foregoing review of motor neuron and muscle fiber properties introduced general features of crustacean neuromuscular systems that govern their response properties and illustrated their

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Harold Atwood diversity. Many of these features are shared with other arthropods, insects in particular. The significant features comprise: 1. Limitation in number of motor neurons per muscle: Excitatory neurons typically range from one to eight, inhibitory neurons from zero to two. Each neuron innervates muscle fibers with many terminals and synapses (i.e., multiterminal innervation). 2. Differentiation of motor neuron properties: Activity patterns differ, giving rise to the phasic-tonic distinction. In addition, synaptic properties of different terminals of a single neuron vary from one muscle fiber to another and are important for muscle fiber recruitment within a muscle. 3. Differentiation of muscle fiber properties: Tremendous variation of structure, biochemical properties, and membrane properties occur between and within muscles. These features impart differences in strength and contraction speed to individual muscle fibers, and to collections of them within muscles. 4. Matching of nerve terminal and muscle fiber properties: Typically, the performance of individual muscle fibers is due to a combined contribution of appropriate synaptic input and muscle fiber response properties. Consideration of individual muscles in functional terms involves discerning how the appropriate nerve and muscle properties have been selected and combined (during evolution) from the large assortment of available response characteristics. In fact, each muscle has its own distinctive features. Phasic and Tonic Neuromuscular Systems The most clear-cut example of phasic and tonic neuromuscular systems is that of the crayfish and lobster abdominal flexor and extensor muscles. Kennedy and Takeda (1965a, 1965b) established the phasic-tonic distinction, showing that most of the abdominal muscle mass in a crayfish is usually inactive and recruited only for escape and vigorous backward swimming, while much smaller superficial slow muscles are active most of the time, regulating postural control and slow movement. The respective neuromuscular properties are appropriate for these activities. Phasic motor neurons, with large output of neurotransmitter per impulse, supply fast-acting, excitable muscle fibers, frequently spiking, that possess short sarcomeres and high myosin ATPase activity, along with other specialized molecular features known to be geared for rapid contraction. Neither synapses nor muscle fibers are able to support sustained activity. Among crustacean muscles, these fast-acting muscles show more uniformity of synaptic and muscle fiber properties than most others. In contrast, the slow abdominal muscles are innervated by several motor axons (six in crayfish) which have varying amounts of neural activity: some are active most of the time, while at least one is active only when abdominal adjustment, rather than postural maintenance, is required. These motor neurons generate relatively small, fatigue-resistant EJPs that produce graded muscle fiber contractions through summation in accordance with the incoming frequency and patterns of motor impulses. The muscle fibers are correspondingly slow and fatigue-resistant, though at least two well-defined fiber types have emerged from detailed studies of the Norway lobster (Holmes et al. 2002). Thus, the very different performances of phasic and tonic neuromuscular systems can be accounted for by matched nerve and muscle fiber properties. Multifunctional Neuromuscular Systems Individual muscles of limbs and other appendages are often able to produce both fast and slow movements, and in some cases postural maintenance, with very few motor neurons. For



Peripheral Components of Crustacean Motor Systems

convenience, such muscles can be termed “multifunctional,” and since the muscles of a limb often share motor neurons, the assemblage of muscles can be referred to as a “multifunctional neuromuscular system.” Many limb muscles have separate regions devoted to different types of movement. As examples, the power-stroke swimming muscles (Fig. 4.5) and the levator of the eyestalk of portunid crabs (Hoyle 1968a) possess separate bands of muscle fibers specialized for maintained activity (pink fibers) and transient rapid activity (white fibers). In the eyestalk muscle, innervation by four motor axons is distributed differentially:  two phasic-type (fast) excitatory motor axons supply both types of muscle fiber, while a tonic (slow) excitor and an inhibitory axon supply only the pink (fatigue-resistant) fibers. This arrangement imparts more refined control over muscle fibers devoted to slow and maintained movement and less refined control over those assigned to rapid, transient movements. Inhibitory innervation to the pink fibers provides for regulation of their tension during slow movement, and also for reduction of their activity during rapid activity. This theme appears in more subtle form in the peripheral leg muscles. Among the more thoroughly studied leg muscles, bands of muscle fibers specialized for slow or fast movement are often present. Examples include the leg closer muscles of several crabs, and the “cutter” claw of the American lobster. However, muscle fiber heterogeneity is often encountered within leg muscles (Fig. 4.2A; Table 4.1). In such cases, the muscle fibers characterized as slow or tonically contracting receive strong input from tonic (“slow”) motor neurons, while fast muscle fibers are selectively driven by phasic (“fast”) motor neurons (Figs. 4.2 and 4.3A; Table 4.1). Inhibitory input is much stronger for slow muscle fibers. Thus, within a muscle, subsets of muscle fibers can be selectively driven by a phasic or a tonic motor neuron, and selectively inhibited. Leg muscles receiving only one motor axon (opener and stretcher muscles in particular) are able to generate fast and slow movements by synaptic recruitment of fast and slow muscle fibers, respectively. Large, slow, summating EJPs are typically produced in slow-acting muscle fibers, leading them to contract at low frequencies of stimulation. At higher frequencies of stimulation, fast muscle fibers are depolarized to the E-C threshold and contract; graded spikes often add to contraction speed (Fig. 4.7B). Strong inhibitory input to slow muscle fibers provides a mechanism for removing their participation during rapid movements. Thus, a single excitatory motor axon can accomplish multiple types of movement, although fast movement may be less powerful than in muscles that have a specialized phasic axon in addition. The roles of specific and common inhibitors have been investigated in several studies through recordings from intact animals. Action of specific inhibitors is essential for independent movement of opener and stretcher muscles. In most cases, specific inhibitors are coactivated with the excitor, and the intensity of inhibition is adjusted in accordance with the locomotory requirements for the target muscle (Wilson and Davis 1965, Spirito 1970). In crayfish, no evidence has been found for control of the timing of inhibitory impulses to take maximal advantage of presynaptic inhibition; instead, this type of inhibition appears to be an “add-on” to the more prevalent postsynaptic inhibition, adding generally to overall effectiveness of inhibition. A reported exception is found in fiddler crabs (Spirito 1970), where selective timing of inhibitory impulses to opener and stretcher muscles provides for optimal inhibition. An additional role for specific inhibitors was suggested by recordings from the crab Cancer magister, in which EJPs appeared to be suppressed by coactivation of inhibition for part of a walking cycle, and suddenly rebounded to large amplitude, suggesting that excitatory facilitation could be generated and maintained during intense inhibition (Atwood and Walcott 1965). Such effects have been demonstrated in studies of single innervated crab muscle fibers (Wiens and Atwood 1975, Atwood and Tse 1993).

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Harold Atwood The role of the common inhibitor, which innervates all leg muscles (Fig. 4.3), was elegantly clarified by Ballantyne and Rathmayer (1981), who recorded axonal activity from intact walking crabs. They found that firing of the common inhibitor speeded relaxation of muscle tension after a burst of activity, enhancing the trough-to-peak transient tension generated by repeated excitatory motor bursts. This effect is attributed to selective inhibition of the slowly contracting muscle fibers, which effectively eliminates their contribution to muscle tension; this contribution, if allowed to persist, reduces overall contraction speed (Atwood 1973, Wiens 1989). Since all leg muscles are affected, though to varying degrees, the overall effect of the common inhibitor is to enhance the speed of movement of the entire leg (with concomitant sacrifice of total force). Matching of Synapse and Muscle Fiber Although the detailed mechanisms responsible for the precise matching of the properties of neuromuscular transmission and muscle fibers are not well understood in crustaceans, recent work in Drosophila has uncovered a complex two-way communication between nerve and muscle that employs several intracellular signaling pathways, and these are likely to be present also, in some form, in crustaceans. At present, we can say that the ultimate properties of synapses formed on different muscle fibers by a single motor axon are heavily dependent on retrograde influences of the muscle fiber, giving rise to the wide diversity of synaptic properties, especially in limb muscles (Frank 1973, see ­chapter 6 in this volume). However, predetermined properties of the motor neuron set the boundaries of synaptic performance, as is evident from the differences between phasic and tonic synapses on single muscle fibers (Fig. 4.3A). Environmental influences also contribute, especially during development. The meticulous research on lateralization of lobster claw muscles resulting in “crusher” and “cutter” claws with different constituent muscle fibers and neuromuscular transmission is due in part to relative neural activity during a critical period in development (review: Govind 1992). Even in postdevelopmental stages, crustacean neuromuscular junctions are malleable:  the phasic neuron of the crayfish claw closer muscle can acquire some tonic structural and physiological characteristics when its activity is increased, and this effect is normally exhibited during seasonal changes in the animal’s activity (review: Lnenicka and Morley 2002). Neurohormonal Modulation Neuromuscular systems are subject to modulation by neurohormones (released by nerve cells into the circulation and acting on neuromuscular junctions and/or muscle fibers remote from the site of release), and by local neuromodulators (materials released at synapses close to their targets but not directly activating fast ionotropic receptors) (see c­ hapter  5 in this volume). In crustaceans, the best-studied neurohormones affecting skeletal muscles are the amines octopamine and serotonin (5-HT), which impose generalized effects on aggressive behavior and escape responses; modulation of neuromuscular systems is one component of their total action. These circulating neurohormones have powerful excitatory effects on both neurotransmission and muscle (see reviews by Beltz and Kravitz 2002 and Atwood and Klose 2009b). Dopamine has a counteracting inhibitory effect on limb muscles. Known neuromodulators (some of which also serve as neurohormones) include peptides such as proctolin, allostatin, and members of the FMRFamide peptide family. In fact, large numbers of related peptides have been discovered in arthropods, and more have been found to date in insects than in crustaceans. Many details of the cellular actions of these various materials have been worked out; however, the circumstances under which they are recruited during



Peripheral Components of Crustacean Motor Systems

locomotor activity to modify peripheral neuromuscular functions are not fully understood, especially for peptides. Amines Serotonin is known to be released from the crustacean pericardial organ, which is well supplied with serotonergic nerve terminals. Serotonin dramatically enhances excitatory neurotransmission in crustacean skeletal muscle. Both motor nerve terminals and muscle fibers are affected: on application of serotonin, the former release more neurotransmitter per nerve impulse, while the latter contract more vigorously and become more excitable with more frequent production of action potentials. The enhancement of transmitter release, which is prolonged, is not linked to a larger influx of Ca 2+ at motor nerve terminals. At the cellular level, activation of the putative presynaptic 5-HT receptors on excitatory nerve terminals potentiates neurotransmitter release through positive enhancement of two second messenger pathways: (1) adenylyl cyclase, leading to cyclic adenosine monophosphate (cAMP) production, which in turn produces downstream effects on exchange factor activated by cAMP (Epac) and hyperpolarization and cyclic nucleotide-activated (HCN) ion channels; these effects lead to prolonged enhancement of neurotransmission (Dixon and Atwood 1989, Beaumont and Zucker 2000, Zhong and Zucker 2005); and (2)  phospholipase C and production of inositol 1,4,5-trisphosphate, thought to generate initial enhancement. Recent reviews on amine modulation have been provided by Harris-Warrick and Johnson (2002) and Beltz and Kravitz (2002). Octopamine, which is also released from the pericardial organ, produces modest enhancement of crustacean neuromuscular transmission and contraction in many limb muscles, largely by direct effects on muscle fibers. In crayfish, there is some enhancement of neurotransmitter output and EJPs, and the degree of enhancement depends on the activity of the target motor neuron. The details of octopamine’s actions, including receptors and second messenger pathways, have been studied more thoroughly in insects (review: Atwood and Klose 2009b). Peptides A large number of bioactive peptides exert overlapping effects on crustacean neuromuscular systems. The pentapeptide proctolin was identified as a neuromodulator for crayfish slow flexor muscles, where it occurs in three of the excitatory motor neurons and promotes contraction by enhancing muscle fiber membrane calcium channel activity (Bishop et al. 1991). More recently, proctolin was found to increase neurotransmitter release at phasic and tonic excitors of a crab closer muscle, through positive modulation of P/Q-type Ca 2+ channel activity (Rathmayer et al. 2002a). In this muscle, a member of the FMRFamide peptide family also modulates Ca2+ channel activity; here, the target is the N-type channel, which pharmacological tests demonstrated only on terminals of the phasic excitor. Hence, in this example, peptidergic modulation would allow differential tuning of the two excitatory motor neurons. An additional case of peptidergic modulation involving proctolin was described in the isopod Idotea, in which proctolin and allostatin serve as positive and negative modulators of neuromuscular function, respectively, through complex pre- and postsynaptic effects (Rathmayer et al. 2002b). A detailed investigation of the neuromuscular enhancing effects of one member of the FMRFamide family, DF2, reveals that its major action on crayfish fast extensor muscles is presynaptic, manifested as increased neurotransmitter release (Mercier et al. 2002). Both cAMP and cyclic guanosine monophosphate (cGMP) are required for the modulatory response; contributions of Ca 2+/calmodulin-dependent protein kinase II (CaMKII) and protein kinase C, respectively, mediate the early and late components of the enhancement. Thus, a complicated

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Harold Atwood intracellular biochemical circuit is activated by this neuropeptide. Once again, the consequences for locomotion and behavior remain to be fully clarified.

COMPARATIVE ASPECTS A full comparative account of muscle and neuromuscular systems was published by Hoyle (1983), and brief updates by Atwood and Klose (2009a, 2009b). Among the arthropods, crustaceans and insects share many features of neuromuscular organization and neuromuscular transmission, including (1)  relatively few excitor axons per muscle (some of which are specialized, as in the case of crustacean phasic and tonic motor neurons); (2) use of “common” inhibitory motor neurons; (3) facilitation of synaptic transmission, and summation of EJPs, as mechanisms to recruit muscle fibers to contract; and (4) graded or all-or-none muscle action potentials. Crustaceans and insects differ in that crustaceans have developed a wider range of muscle fiber properties in most locomotory muscles, exploiting all variables very fully. Insects have developed distinctive specialized “synchronous” and “asynchronous” flight muscles and the asynchronous tymbal muscles responsible for sound production in cicadas that are quite different from the normal locomotory muscles controlling limb movements. Both insects and crustaceans differ from other arthropods in having more complex neuromuscular organization of individual motor axons, and in general, fewer motor axons per muscle. However, other arthropods including the major chelicerate groups have been much less intensively investigated. In comparing crustaceans with well-studied examples of the other “major” phyla (particularly chordates, molluscs, and nematodes), one can observe that the theme of polyneuronal, multiterminal innervation of muscle fibers is often preserved among invertebrate phyla, whereas in skeletal muscles of the more advanced vertebrates, innervation of skeletal muscle fibers at a single end-plate is more often seen. In muscles of lower vertebrates, slow or tonic muscle fibers with multiterminal innervation are frequently found, but these are rare in mammals. In typical skeletal muscles, excitation at the end-plate by the motor axon’s action potential typically generates a propagated muscle action potential and a twitch contraction. Summation of twitch contractions in response to repeated activation of the innervating motor neuron can produce variable amounts of tension, between twitch and tetanus, in a single muscle fiber. Low-level, long-lasting contractions of individual muscle fibers, typical of many crustacean fibers, are not a feature of mammalian twitch muscle fibers, but are utilized in tonus fibers of frogs and other lower vertebrates. Gradation of muscle contraction in most vertebrate skeletal muscles also involves recruitment of motor units (small groups of muscle fibers innervated by individual motor neurons), and each muscle typically has numerous motor units to allow for optimal gradation in strength of contraction. Fast and slow contractions are achieved in part by specialization of muscle fiber properties, although the range of variation of muscle fiber properties is less than in crustaceans. In molluscs, muscles are innervated by varying numbers of motor neurons, many of which are cholinergic and a few inhibitory. Modulation by neuropeptides, some of which are likely cotransmitters, appears to be more important for molluscan muscles than in crustaceans. Fast and slow contractions are achieved in some molluscs through specialization of muscle fiber properties and by recruitment of appropriate muscle fiber types via innervating motor neurons, as in the swimming pteropod mollusc Clione. An extreme example of tonic muscle specialization is provided by the byssal retractors of sessile mussels such as Mytilus, and extreme strength is characteristic of adductor muscles of bivalve molluscs. Thus, in major phyla, various neuromuscular mechanisms have been developed to achieve the goals of gradation of strength,



Peripheral Components of Crustacean Motor Systems

variable speed of locomotion, tonic contraction, and provision of separate systems for fast and slow actions (Hoyle 1983).

FUTURE DIRECTIONS Crustacean neuromuscular systems afford excellent opportunities for investigation of fundamental aspects of synaptic function and nerve-muscle interaction. The large size, unusual synaptic properties, and unique identification of neural components coupled with the large size and wide range of properties of muscular components is experimentally attractive and these features continue to offer a wide field for exploration. An example of a problem that could be further explored to advantage in crustacean muscle fibers is the basis for the range in E-C threshold in different muscle fibers. During the last two decades, crustacean neuromuscular systems have been eclipsed in the neuroscience arena by the rapid development of experimentation in the fruit fly Drosophila, in which the larval neuromuscular system has proven to be amenable to neurophysiological experimentation, besides offering the advantages of modern genetic information and sophisticated genetic techniques. In several studies, genetic information and specific proteins found in Drosophila have stimulated work on crustacean counterparts as a prelude to answering questions about synaptic differentiation in a physiologically favorable system (e.g., Jeromin et al. 1999). As more genetic information and cloned genes become available from crustacean species, questions that depend on such information for their resolution will be open to investigation. Modern techniques for commanding and timing protein expression in selected locations could be exploited in crustaceans, as they presently are in Drosophila. Among the topics that can be advantageously tackled in crustaceans are: the mechanisms of long-term synaptic adaptation in response to activity patterns, adaptation of neuromuscular systems to acute and chronic environmental conditions, the mechanisms for matching of muscle fiber and synaptic properties, muscle fiber differentiation, and mechanisms underlying synaptic facilitation and depression. In addition to pursuit of mechanisms at the cellular and subcellular levels, questions about the roles of these features in behavior will continue to be explored as an ongoing endeavor.

SUMMARY AND CONCLUSIONS Crustacean neuromuscular systems have evolved with relatively few peripheral neural components: most muscles are supplied by only a few excitatory motor neurons and many receive in addition one or two inhibitory motor neurons. In strength of contraction and range of speed and precision, crustacean muscles equal and often exceed what can be achieved in muscles of other advanced phyla. The features that contribute to this exceptional versatility are apparent in both the motor neurons and their target muscle fibers. On the neural side, synaptic properties such as initial release of neurotransmitters and facilitation of transmitter release cover a wide range and differ strikingly from neuron to neuron and also at different endings of a single neuron. These synaptic features contribute to selective recruitment of groups of muscle fibers for specific actions. Matching this feature, the muscle fibers are differentiated into a variety of functional types, specialized for fast or slow contraction, and for brief or sustained activity, with many intermediate forms. Molecular and structural variation is more diverse in crustacean muscle fibers than in those of other major taxonomic groups. Matching of nerve and muscle properties is a striking feature in crustacean muscles that is set up during development to produce a refined system optimally adapted for the needs of the species.

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ACKNOWLEDGMENTS Research on crustacean neuromuscular systems in the author’s laboratory was supported since 1965 by the Natural Science and Engineering Research Council of Canada, and since 1981 by the Medical Research Council of Canada (now Canadian Institutes of Health Research). The author gratefully acknowledges the past help and direction of his mentors in crustacean neurobiology, Graham Hoyle and C.A.G. (Kees) Wiersma; and the outstanding contributions of two former laboratory members whose work was prematurely terminated: Fred Lang and C.K. Govind.

REFERENCES Atwood, H.L. 1972. Crustacean muscle. Pages 421–489 in G.H. Bourne, editor. The structure and function of muscle. Academic Press, New York. Atwood, H.L. 1973. Crustacean motor units. Pages 87–104 in R.B. Stein, K.G. Pearson, R.S. Smith, and J.B. Redford, editors. Control of posture and locomotion. Plenum Press, New York. Atwood, H.L. 1976. Organization and synaptic physiology of crustacean neuromuscular systems. Pages 291–391 in G.A. Kerkut and J.W. Phillis, editors. Progress in neurobiology, Vol. 7. Pergamon Press, Oxford, U.K. Atwood, H.L. 1982. Synapses and neurotransmitters. Pages 105–150 in H.L. Atwood and D.C. Sandeman, editors. The biology of Crustacea, Vol. 3, Neurobiology: structure and function. Academic Press, New York. Atwood, H.L., and G.D. Bittner. 1971. Matching of excitatory and inhibitory inputs to crustacean muscle fibers. Journal of Neurophysiology 34:157–170. Atwood, H.L., and S. Karunanithi. 2002. Diversification of synaptic strength: presynaptic elements. Nature Reviews Neuroscience 3:497–516. Atwood, H.L., and M.K. Klose. 2009a. Comparative biology of invertebrate neuromuscular junctions. Pages 1185–1209 in L.R. Squire, editor. Encyclopedia of neuroscience, Vol. 2. Academic Press, Oxford, U.K. Atwood, H.L., and M.K. Klose. 2009b. Neuromuscular transmission modulation at invertebrate neuromuscular junctions. Pages 671–690 in L.R. Squire, editor. Encyclopedia of neuroscience, Vol. 6. Academic Press, Oxford, U.K. Atwood, H.L., and F.W. Tse. 1993. Physiological aspects of presynaptic inhibition. Pages 19–65 in S.K. Malhotra, editor. Advances in neural science, Vol. 1. JAI Press, Greenwich, CT. Atwood, H.L., and B. Walcott. 1965. Recording of electrical activity and movement from legs of walking crabs. Canadian Journal of Zoology 43:657–665. Atwood, H.L., and J.M. Wojtowicz. 1986. Short-term and long-term plasticity and physiological differentiation of crustacean motor synapses. Pages 275–362 in J.R. Smythies and R.J. Bradley, editors. International review of neurobiology, Vol. 28. Academic Press, Orlando. Atwood, H.L., and J.M. Wojtowicz. 1999. Silent synapses in neural plasticity: current evidence. Learning and Memory 6:542–571. Atwood, H.L., G. Hoyle, and T. Smyth. 1965. Mechanical and electrical responses of single innervated crab-muscle fibres. Journal of Physiology (London) 180:449–482. Ballantyne, D., and W. Rathmayer. 1981. On the function of the common inhibitory neurone in the walking legs of the crab, Eriphia spinifrons. Journal of Comparative Physiology A 143:111–122. Beaumont, V., and R.S. Zucker. 2000. Enhancement of synaptic transmission by cyclic AMP modulation of presynaptic Ih channels. Nature Neuroscience 3:133–141. Beltz, B.S., and E.A. Kravitz. 2002. Serotonin in crustacean systems: more than a half century of fundamental discoveries. Pages 141–163 in K. Wiese, editor. Crustacean experimental systems in neurobiology. Springer-Verlag, Berlin.



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Bishop, C.A., M.E. Krouse, and J.J. Wine. 1991. Peptide cotransmitter potentiates calcium channel activity in crayfish skeletal muscle. Journal of Neuroscience 11:269–276. Bliss, D.E., editor-in-chief. 1982. The biology of Crustacea, Vols. 3 and 4. Academic Press, New York. Bradacs, H., R.L. Cooper, M. Msghina, and H.L. Atwood. 1997. Differential physiology and morphology of phasic and tonic motor axons in a crayfish limb extensor muscle. Journal of Experimental Biology 200:677–691. Chapple, W.D. 1982. Muscle. Pages 151–184 in H.L. Atwood and D.C. Sandeman,, editors. The biology of Crustacea, Vol. 3, Neurobiology: structure and function. Academic Press, New York. Cooper, R.L., C.C. Harrington, L. Marin, and H.L. Atwood. 1996. Quantal release at visualized terminals of a crayfish motor axon: intraterminal and regional differences. Journal of Comparative Neurology 375:583–600. Dixon, D.B., and H.L. Atwood. 1989. Conjoint action of phosphatidylinositol and adenylate cyclase systems in serotonin-induced facilitation at the crayfish neuromuscular junction. Journal of Neurophysiology 62:1252–1259. Dudel, J., and S.W. Kuffler. 1961. Presynaptic inhibition at the crayfish neuromuscular junction. Journal of Physiology (London) 155: 543–562. Frank, E. 1973. Matching of facilitation at the neuromuscular junction of the lobster: a possible case for the influence of muscle on nerve. Journal of Physiology (London) 233:635–658. Franzini-Armstrong, C., A.B. Eastwood, and L.D. Peachey. 1986. Shape and disposition of clefts, tubules, and sarcoplasmic reticulum in long and short sarcomere fibers of crab and crayfish. Cell and Tissue Research 244:9–19. Goblet, C., and Y. Mounier. 1986. Calcium-induced calcium release from the sarcoplasmic reticulum in skinned crab muscle fibres. Cell Calcium 7:61–72. Govind, C.K. 1992. Claw asymmetry in lobsters: case study in developmental neuroethology. Journal of Neurobiology 23:1423–1445. Govind, C.K., and H.L. Atwood. 1982. Organization of neuromuscular systems. Pages 63–103 in H.L. Atwood and D.C. Sandeman, editors. The biology of Crustacea, Vol. 3, Neurobiology: structure and function. Academic Press, New York. Harris-Warrick, R.M., and B.R. Johnson. 2002. Cellular and molecular mechanisms of amine modulation in Crustacea. Pages 23–43 in K. Wiese, editor. Crustacean experimental systems in neurobiology. Springer-Verlag, Berlin, Germany. Holmes, J.M., D.M. Neil, S. Galler, and K. Hilber. 2002. Correlation of the synaptic and mechanical properties of two slow fibre phenotypes in a crustacean muscle. Pages 292–304 in K. Wiese, editor. The crustacean nervous system. Springer-Verlag, Berlin, Germany. Hoyle, G. 1968a. Correlated physiological and ultrastructural studies on specialised muscles. Ia. Neuromuscular physiology of the levator of the eyestalk of Podophthalmus vigil (Weber). Journal of Experimental Zoology 167:471–486. Hoyle, G. 1968b. Resting tension, “negative” contraction and “break” contraction in specialised crustacean muscle fibers. Journal of Experimental Zoology 167:551–566. Hoyle, G. 1983. Muscles and their neural control. John Wiley & Sons, New York. Hoyle, G., and C.A.G. Wiersma. 1958. Excitation at neuromuscular junctions in Crustacea. Journal of Physiology (London) 143:403–425. Jahromi, S.S., and H.L. Atwood. 1971. Structural and contractile properties of lobster leg-muscle fibers. Journal of Experimental Zoology 176:475–486. Jeromin, A., A.J. Shayan, M. Msghina, J. Roder, and H.L. Atwood. 1999. Crustacean frequenins: molecular cloning and differential location at neuromuscular junctions. Journal of Neurobiology 41:165–175. Kennedy, D., and K. Takeda. 1965a. Reflex control of abdominal flexor muscles in the crayfish. I. The twitch system. Journal of Experimental Biology 43:11–227. Kennedy, D., and K. Takeda. 1965b. Reflex control of abdominal flexor muscles in the crayfish. II. The tonic system. Journal of Experimental Biology 43:229–246. Lnenicka, G.M., and E.J. Morley. 2002. Activity-dependent development and plasticity of crustacean motor terminals. Pages 266–281 in K. Wiese, editor. The crustacean nervous system. Springer-Verlag, Berlin, Germany.

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Harold Atwood Mendelson, M. 1969. Electrical and mechanical characteristics of a very fast lobster muscle. Journal of Cell Biology 42:548–563. Mercier, A.J., A. Badhwar, A.D. Weston, and M. Klose. 2002. Intracellular signals that mediate synaptic modulation by a FMRFamide-like neuropeptide in crayfish. Pages 49–62 in K. Wiese, editor. The crustacean nervous system. Springer-Verlag, Berlin, Germany. Millar, A.G., R.S. Zucker, G.C. Ellis-Davies, M.P. Charlton, and H.L. Atwood. 2005. Calcium sensitivity of neurotransmitter release differs at phasic and tonic synapses. Journal of Neuroscience 25:3113–3125. Miwa, A., M. Ui, and N. Kawai. 1990. G-protein is coupled to presynaptic glutamate and GABA receptors in lobster neuromuscular synapse. Journal of Neurophysiology 63:173–180. Msghina, M., A.G. Millar, M.P. Charlton, C.K. Govind, and H.L. Atwood. 1999. Calcium entry related to active zones and differences in transmitter release at phasic and tonic synapses. Journal of Neuroscience 19:8419–8434. Mykles, D.L. 1985. Heterogeneity of myofibrillar proteins in lobster fast and slow muscles: variants of troponin, paramyosin, and myosin light chains comprise four distinct protein assemblages. Journal of Experimental Zoology 234:23–32. Nguyen, P.V., L. Marin, and H.L. Atwood. 1997. Synaptic physiology and mitochondrial function in crayfish tonic and phasic motor neurons. Journal of Neurophysiology 78:281–294. Orkand, R.K. 1962a. The relation between membrane potential and contraction in single muscle fibres. Journal of Physiology (London) 161:143–159. Orkand, R.K. 1962b. Chemical inhibition of contraction in directly stimulated crayfish muscle fibres. Journal of Physiology (London) 164:103–115. Pan, B., and R.S. Zucker. 2009. A general model of synaptic transmission and short-term plasticity. Neuron 62:539–554. Peled, E.S., and E.Y. Isacoff. 2011. Optical quantal analysis of synaptic transmission in wild-type and rab3-mutant Drosophila motor axons. Nature Neuroscience 14:519–526. Quigley, P.A., M. Msghina, C.K. Govind, and H.L. Atwood. 1999. Visible evidence for differences in synaptic effectiveness with activity-dependent vesicular uptake and release of FM1-43. Journal of Neurophysiology 81:356–370. Rathmayer, W., and S. Djokaj. 2000. Presynaptic inhibition and the participation of GABAB receptors at neuromuscular junctions of the crab Eriphia spinifrons. Journal of Comparative Physiology A 186:287–298. Rathmayer, W., and C. Erxleben. 1983. Identified muscle fibers in a crab. I. Characteristics of excitatory and inhibitory neuromuscular transmission. Journal of Comparative Physiology A 152:411–420. Rathmayer, W., and M. Hammelsbeck. 1985. Identified muscle fibres in a crab. Differences in facilitation properties. Journal of Experimental Biology 116:291–300. Rathmayer, W., and L. Maier. 1987. Muscle fiber types in crabs: studies on single identified muscle fibers. American Zoologist 27:1067–1077. Rathmayer, W., S. Djokaj, A. Gaydukov, and S. Kreissl. 2002a. The neuromuscular junctions of the slow and the fast excitatory axon in the closer of the crab Eriphia spinifrons are endowed with different Ca2+ channel types and allow neuron-specific modulation of transmitter release by two neuropeptides. Journal of Neuroscience 22:708–717. Rathmayer, W., C. Erxleben, S. Djokaj, A. Gaydukov, S. Kreissl, and T. Weiss. 2002b. Antagonistic modulation of neuromuscular parameters in crustaceans by the peptides proctolin and allatostatin, contained in identified motor neurons. Pages 2–19 in K. Wiese, editor. The crustacean nervous system. Springer-Verlag, Berlin, Germany. Rojas, E., V. Nassar-Gentina, M.E. Pollard, and M. Luxoro. 1992. Mechanisms of calcium release from terminal cisternae in crustacean muscle. Pages 305–317 in G.B. Frank, C.P. Bianchi, and H.E.D.J. ter Keurs, editors. Excitation-contraction coupling in skeletal, cardiac, and smooth muscle. Advances in experimental medicine and biology, Vol. 311. Plenum Press, New York. Rosenbluth, J. 1969. Sarcoplasmic reticulum of an unusually fast-acting crustacean muscle. Journal of Cell Biology 48:174–188.



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Silverman, H., and M.P. Charlton. 1980. A fast-oxidative crustacean muscle: histochemical comparison with other crustacean muscle. Journal of Experimental Zoology 211:267–273. Silverman, H., W.J. Costello, and D.L. Mykles. 1987. Morphological fiber type correlates of physiological and biochemical properties in crustacean muscle. American Zoologist 27:1011–1019. Silverman-Gavrila, L.B., P.M.R. Orth, and M.P. Charlton. 2005. Phosphorylation-dependent low-frequency depression at phasic synapses of a crayfish motoneuron. Journal of Neuroscience 25:3186–3180. Spirito, C.P. 1970. Reflex control of the opener and stretcher muscles in the cheliped of the fiddler crab, Uca pugnax. Zeitschrift für Vergleichende Physiologie 68:211–228. Taylor, G.M. 2000. Maximum force production: why are crabs so strong? Proceedings of the Royal Society of London B 267:1475–1480. Tse, F.W., C.K. Govind, and H.L. Atwood. 1983. Diverse fiber composition of swimming muscles in the blue crab, Callinectes sapidus. Canadian Journal of Zoology 61:52–59. Ushio, H., and S. Watabe. 1993. Crayfish skeletal muscle requires both influx of external Ca 2+ and Ca 2+ release from internal stores for contraction. Journal of Experimental Biology 181:95–105. Wiens, T.J. 1989. Common and specific inhibition in leg muscles of decapods: sharpened distinctions. Journal of Neurobiology 20:458–469. Wiens, T.J., and H.L. Atwood. 1975. Dual inhibitory control in crab leg muscles. Journal of Comparative Physiology 99:211–230. Wiens, T.J., and W. Rathmayer. 1985. The distribution of the common inhibitory neuron in brachyuran limb musculature. I. Target muscles. Journal of Comparative Physiology A 156:305–313. Wiens, T.J., J. Pearce, and C. K. Govind. 1991. Neuromuscular properties of the quintuply innervated flexor muscle in lobster limbs. Canadian Journal of Zoology 69:477–488. Wiersma, C.A.G. 1951. A bifunctional single motor axon system of a crustacean muscle. Journal of Experimental Biology 28:13–21. Wiersma, C.A.G. 1961. The neuromuscular system. Pages 191–240 in T.H. Waterman, editor. The physiology of Crustacea, Vol. 2, Sense organs, integration, and behavior. Academic Press, New York. Wiese, K., editor. 2002a. The crustacean nervous system. Springer-Verlag, Berlin, Germany. Wiese, K., editor. 2002b. Crustacean experimental systems in neurobiology. Springer-Verlag, Berlin, Germany. Wiese, K., W.-D. Krenz, J. Tautz, H. Reichert, and B. Mulloney, editors. 1990. Frontiers in crustacean neurobiology. Birkhäuser Verlag, Basel, Switzerland. Wilson, D.M., and Davis, W.J. 1965. Nerve impulse patterns and reflex control in the motor system of the crayfish claw. Journal of Experimental Biology 43:193–210. Wojtowicz, J.M., L. Marin, and H.L. Atwood. 1994. Activity-induced changes in synaptic release sites at the crayfish neuromuscular junction. Journal of Neuroscience 14:3688–3702. Zhong, N., and R.S. Zucker. 2005. cAMP acts on exchange protein activated by cAMP/cAMP-regulated guanine nucleotide exchange protein to regulate transmitter release at the crayfish neuromuscular junction. Journal of Neuroscience 25:208–214.

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5 MODULATION OF CRUSTACEAN NETWORKS FOR BEHAVIOR

Ronald Harris-Warrick

Abstract Crustacean neuroscience has played a leading role in studying the mechanisms by which neuromodulators alter simple behaviors. Many small transmitters and peptides modulate crustacean behavioral networks. They can act as synaptically released transmitters, locally released paracrine hormones, or globally released circulating neurohormones, acting over different time courses and concentration ranges. Most modulatory neurons release multiple transmitters, with complex interactions. The actions of these modulators are best understood in three crustacean networks: the heartbeat network in the cardiac ganglion, the swimmeret networks in abdominal ganglia, and rhythmic foregut digestive networks in the stomatogastric ganglion. Modulatory inputs can activate, terminate, or modify the properties of the network’s motor pattern. They can fuse multiple networks together to generate complex, multicomponent movements. Neuromodulators also modify the properties of peripheral muscles and sensory receptors, which must be coordinated with their central effects on behavioral networks. Every component of a neural network is subject to neuromodulation. Neuromodulators alter the intrinsic firing properties of the component neurons, such as rhythmic bursting, bistability, postinhibitory rebound, and spike frequency regulation, by altering the properties of ionic currents that shape neuronal activity. They also modify the strengths of synapses in the networks by pre- and postsynaptic mechanisms, quantitatively rewiring the network. The effects of neuromodulators on a behavior can only be understood from the sum of their effects on the network and its peripheral apparatus. Many major questions remain in understanding how behavioral network modulation occurs, which will continue to be studied in crustacean systems.

INTRODUCTION When The Biology of Crustacea was published almost 30 years ago, the concept of neuromodulation was just starting to be addressed. One of the earliest definitions of a neuromodulator 114



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was given by Ernst Florey (Florey 1967)  based on his studies of neurohormonal actions of amines in Crustacea: “I would like to use the term ‘modulator substance’ for any compound of cellular and nonsynaptic origin that affects the excitability of nerve cells. . . Such modulator substances can affect the responsiveness of nerve cells to transsynaptic actions of presynaptic neurons and they can alter the tendency to spontaneous activity.” This study emphasized hormonal actions of modulators, but it soon became clear that similar actions could arise from synaptically released neuromodulators. In an early major review of the topic, Kupfermann (1979) defined a new class of synaptic events:  “Unlike conventional transmitters, they do not simply excite or inhibit an electrically excitable cell, but rather are involved in altering the effects of other events occurring at the cell.” In 1999, Katz updated the concept of neuromodulation, describing the myriad mechanisms by which neurons communicate beyond simple synaptic transmission. These mechanisms were so diverse, that he defined modulatory actions by the negative: “Any communication between neurons, caused by release of a chemical, that is either not fast, or not point-to-point, or not simply excitation or inhibition will be classified as neuromodulatory” (Katz 1999). It is now clear that there is a continuum of transmitter actions, ranging from classical fast synapses typically using ionotropic receptors, to slow synapses typically using metabotropic receptors, to paracrine and hormonal responses that can last for hours. The major characteristics of a modulatory synapse are: (1) they tend to be slow (taking many msec to begin and lasting anywhere from seconds to hours); (2) their actions tend to be subtle, often not manifesting themselves until the target neuron’s voltage is changed or it receives another synaptic input; (3) they can change the basic integrative properties of a neuron or its intrinsic electrophysiological personality; and (4) they typically act by intracellular biochemical mechanisms involving second messenger signal transduction (Harris-Warrick 1988). However, there are exceptions to each of these rules. Crustacean nervous systems have long served as major models for understanding both the mechanisms and the consequences of neuromodulation in nervous system function. Some of the earliest work on the cellular mechanisms of neuromodulation was performed using the neuromuscular junction on the walking legs, heart, and gut of lobsters and crayfish (Grundfest and Reuben 1961, Florey 1967, Kravitz et al. 1980). Network neuromodulation was early studied in the lobster stomatogastric ganglion (STG; Beltz et al. 1984), cardiac ganglion (CG; Cooke 1966), and postural systems (Livingstone et al. 1980), and the crayfish swimmeret system (Mulloney et  al. 1987). Studies of the mechanisms of behavioral modulation have also used crustaceans (Livingstone et al. 1980, Yeh et al. 1996). In this chapter, I will not give a historical record of research on neuromodulation in Crustacea. Rather, I will summarize the general principles that explain how neuromodulators shape behavior by their actions on the neurons and synapses that form behavioral networks in the central nervous system (CNS). Neuromodulators affect behavior by actions at many levels in the nervous system. In the CNS, they play major roles to switch on or off and modify the output of the neural networks that generate behavior. These networks are anatomically defined in terms of their component neurons and synaptic interconnections, but should be considered as libraries of possibilities, since neuromodulators can affect their output in at least three ways: (1) They can determine which of the component neurons actively participate in network function at any moment. (2) They can alter the intrinsic baseline firing properties and pattern of activity of network neurons. (3) They can alter the strength and dynamics of the synaptic interactions or the functional wiring diagram of the network (Harris-Warrick and Flamm 1986). In many cases, modulatory input is essential for a network to function at all to generate a behaviorally relevant output. When the excitability of motor neurons is modulated, the gain of the drive to the muscles can be changed, and in some cases dramatically reorganized when the motor neurons themselves

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Ronald Harris-Warrick develop rhythmic bursting or bistable properties. At the neuromuscular junction, the synaptic drive from motor neurons to the muscles can be gated up and down by both presynaptic changes in transmitter release and postsynaptic changes in responsiveness. Direct effects of modulators on muscle and muscle contractile properties can also dramatically alter behavior. Sensory feedback, which adjusts the motor pattern to the environmental conditions, and can provide essential input to shape network output, is also subject to neuromodulation. Finally, the modulatory neurons themselves can interact in complex networks, shaping the pattern of modulatory input to a network. In this chapter, I will focus primarily on network modulation. Modulation at the neuromuscular junction and direct modulatory effects on muscle are discussed by Atwood in ­chapter 4 of this volume.

IDENTIFICATION OF NEUROMODULATOR COMPOUNDS IN CRUSTACEA Neuromodulators reach their targets through several paths. Some are released from nerve terminals of modulatory neurons, where they bind directly to pre- or postsynaptic receptors to modify synaptic transmission. Others are released from sites that do not have traditional synaptic architecture. The modulators diffuse to act on nearby neurons, in a mechanism called paracrine hormone function or volume transmission. Still others are released from neurosecretory sites into the hemolymph, acting as circulating hormones to interact with receptors over the entire animal. The major difference between these actions is the effective concentration of modulator at its receptor, which can vary by several orders of magnitude between synaptic and hormonal actions. The mode of delivery of a modulator varies between species. For example, serotonin (5HT) is released into the crab (Cancer irroratus) and American lobster (Homarus americanus) STG from nerve terminals of the sensorymodulatory gastropyloric receptor (GPR) neurons, with a threshold for activation around 10 -6 M. In the spiny lobster Panulirus interruptus, 5HT is not synthesized in GPR neurons, and arrives instead as a hemolymph-borne hormone with threshold activation around 10 -9 M (Beltz et al. 1984, Katz 1989). Given the very broad definition of modulatory actions, it is not surprising that virtually any neurotransmitter can exert a modulatory effect, provided that it interacts with a receptor capable of modifying the intracellular biochemistry. A number of small transmitters with modulatory actions have been identified in crustacean tissues. Acetylcholine is the major transmitter of sensory neurons in crustaceans, and can have both rapid nicotinic responses and muscarinic modulatory effects in the STG of lobsters (Marder 1974, Nagy and Dickinson 1983) and crabs (Dickinson et al. 2008). Glutamate is the fast transmitter of motor neurons, but it can have slow modulatory effects mediated by metabotropic glutamate receptors, in the STG (Krenz et  al. 2000). Histamine is present in the identified inferior ventricular nerve (IVN) interneurons that directly inhibit but indirectly activate lobster STG networks (Claiborne and Selverston 1984, Christie et al. 2004). Histamine also activates a chloride conductance to inhibit neurons in the crayfish X-organ sinus gland (Cebada and García 2007) and CG (Hashemzadeh-Gargari and Freschi 1992). It is a transmitter of arthropod photoreceptors (Stuart et al. 2007) and is involved in presynaptic inhibition of sensory afferents in the coxobasal chordotonal organ in crayfish legs (el Manira and Clarac 1994). A great deal of work has been done on the modulatory effects of the biogenic amines 5HT, dopamine (DA), and octopamine (Kravitz et al. 1980). In arthropods, the functions of norepinephrine are largely replaced by its phenol analog, octopamine, though low concentrations of norepinephrine are found in the lobster CG (Ocorr and Berlind 1983) and prawn ganglia (Hsieh et al. 2006), where it can help to regulate hyperglycemic responses to cold shock. The



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modulatory effects of 5HT, DA, and octopamine at the peripheral neuromuscular junction (see ­chapter 4 in this volume) and sensory responses by the stretch receptor organ (Pasztor 1989)  have been investigated. They are also major modulators of central pattern generator networks (see below). Peptides provide an ever-increasing family of modulators in Crustacea. Christie et al. (2010a) provide an excellent summary of the known crustacean peptides. Peptides were initially identified by biochemical isolation from a large pool of tissue, immunocytochemical localization, or gene expression methods, but recent mass spectrometry (MS)-based methods developed primarily by the Li and Sweedler groups (including MALDI-TOF, MALDI-FTMS, and Tandem MS/MS) have greatly increased the number of identified crustacean neuropeptides. Peptides can now be localized by methods that collect MS spectra from specific sites in slices of tissue, or even whole ganglia (Chen et al. 2010a). It is also possible to quantify the amount of peptide and to measure changes in concentration after behavioral modifications such as feeding or environmental stress (Chen et al. 2010b). To date, hundreds of individual peptides in over 30 families, some with over 20 members, have been localized in crustacean nervous tissue (Christie et al. 2010a). For the large majority of these, their physiological functions have not been studied in any detail.

IDENTIFICATION OF RECEPTORS FOR NEUROMODULATORS As described above, nearly any neurotransmitter can act as a neuromodulator if it interacts with the appropriate receptor. Accordingly, identification and localization of receptors for these compounds in Crustacea will tell us much about their function. No crustacean neuropeptide receptors have been cloned yet, but a number of monoamine receptors are now cloned. An octopamine/tyramine receptor has been cloned from the freshwater prawn Macrobrachium rosenbergii (Reyes-Colón et al. 2010). It is not certain whether tyramine (the precursor of octopamine) functions as a neuromodulator in Crustacea, though some of its physiological actions are different from octopamine in Drosophila (Selcho et  al. 2012). This receptor is widely expressed in prawn central ganglia, consistent with a role for octopamine in the control of posture (Harris-Warrick and Kravitz 1984), swimmeret control (Tschuluun et al. 2009), and regulation of aggression and dominance status (Pedetta et al. 2010). Two 5HT receptors have been cloned from the spiny lobster P. interruptus (Clark et al. 2004), the crayfish Procambarus clarkii (Sosa et al. 2004, Spitzer et al. 2008), and the prawn M. rosenbergii (Vázquez-Acevedo et  al. 2009); they show strong sequence conservation, especially in the central functional core of the protein. The 5HT1αPan receptor is negatively coupled to cyclic adenosine monophosphate (cAMP), while the 5HT2βPan receptor activates a phospholipase C pathway, with phosphatidylinositidol 4,5-bisphosphate (PIP 2) hydrolysis yielding elevations in DAG (diacylglycerol) and IP3. Interestingly, the P. interruptus 5HT2 receptor shows significant constitutive activity even in the absence of 5HT, due to replacement of a single amino acid in the third transmembrane domain (Clark et al. 2004). This suggests that the receptor’s Gq metabolic pathway has significant constitutive activity even when modulatory inputs are silent. As expected of this transmitter with its many effects on behavior, the receptors are widely distributed throughout the CNS and STG. Similarly, Baro’s group has cloned two D1-type and one D2-type DA receptors from P. interruptus (Clark and Baro 2006, 2007). In the lobster STG, the three DA receptors are localized exclusively in the fine neuropil, suggesting an important role in regulation of synaptic transmission (Clark et al. 2008). Studies with receptor-specific antibody antagonists suggest that in lobster membranes, DA1αPan couples with Gs and Gq, while DA1βPan couples with GS exclusively,

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Ronald Harris-Warrick and DA 2αPan couples to Gi exclusively to reduce cAMP levels (Clark et al. 2008). However, in heterologous HEK273 cells, these receptors can activate additional G proteins (Clark and Baro 2007). These careful studies highlight the caution that must be taken in interpreting results from heterologous cell lines.

MODULATION OF SPECIFIC CRUSTACEAN NEURAL NETWORKS To understand how neuromodulators sculpt behavior, it is necessary to understand their effects on the neural networks that generate the timing, phasing, and intensity cues that shape the behavior. Detailed studies are only possible in a few systems, where the neuronal composition and synaptic interactions of the networks are sufficiently worked out to identify modulator sites of action. The great majority of these studies have been carried out on the central pattern generator (CPG) networks in the STG, which drive rhythmic behaviors in the crustacean foregut. Less detailed studies have been performed in the CG, which controls heartbeat, and in the abdominal networks that control swimmeret movement. Additional systems are described in other chapters, including modulatory control of the escape system (see ­chapter 15 in this volume), and of the postural and walking systems (see ­chapter 14 in this volume).

MODULATION IN THE CARDIAC GANGLION AND THE HEARTBEAT CONTROL SYSTEM The CG is a branching nerve trunk containing from 6 to 16 neurons that provides rhythmic neural drive to the crustacean heart muscle (Cooke 2002) (Fig. 5.1A). In most crustaceans, the heart is neurogenic: rhythmic drive from the CG determines the contraction frequency. When dissected from the heart, the isolated CG still generates a rhythmic output, showing that it is an autonomous oscillator network. In decapods, the CG usually contains nine neurons, of which four are “small cells” that have intrinsic oscillatory capability and are considered the pacemaker neurons of the network. They provide the rhythmic synaptic drive to the five “large cells,” which are motor neurons innervating the muscle. However, all the neurons are capable of generating bursts of action potentials, due to the existence of a large somatic calcium-dependent “driver potential.” A number of neurohormones were first detected by their cardioacceleratory (chronotropic) effects or their ability to increase cardiac contraction strength (inotropic effects). As the CG sits inside the heart, it is accessible to all hormones released from the pericardial organ (PO), which is a major source of neurohormones in Crustacea (Fu et  al. 2005, Christie et  al. 2010a). The monoamines 5HT, DA, and octopamine have varied effects on CG activity (Cruz-Bermúdez and Marder 2007). For example, 5HT and DA increase cycle frequency and strength of motor neuron bursts (Fig. 5.1C,D); the increase in cycle frequency arises primarily from direct depolarizing effects on the small cells (Florey and Rathmayer 1978). Octopamine, in contrast, has variable effects depending on the species:  in the crayfish Astacus leptodactylus and the crab Eriphia spinifrons, octopamine exerts a biphasic effect, initially decreasing the cycle frequency before powerfully accelerating it (Florey and Rathmayer 1978), while in Portunus it decreases the burst frequency while increasing the burst duration; the CG motor neurons are hyperpolarized (Benson and Cooke 1984). In the crab Cancer borealis, octopamine and histamine have little or no effect on the isolated CG (Cruz-Bermúdez and Marder 2007). In H. americanus, octopamine evokes a significant elevation of cAMP in the heart, probably reflecting a direct effect on the muscle (Goy 2005).

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Fig. 5.1. Central-peripheral matching of dopamine (DA) modulation in the cardiac ganglion (CG). (A) Picture of the CG, showing the locations of the large cells (above) and small cells (below) (from Cooke 2002, with permission from Marine Biological Laboratory, Woods Hole, MA). (B) Tyrosine hydroxylaselike immunoreactivity in the CG of Callinectes sapidus. A single tyrosine hydroxylase-like immunoreactive fiber reaches the CG by each dorsal nerve (indicated by arrows). The fibers branch and terminate in the region of three large motor neurons labeled with neurobiotin (indicated by asterisks). Scale bar: 100 µm. (C) Effect of DA to strengthen muscle contractions and accelerate the cardiac cycle in the semi-intact whole heart-CG preparation. C1: top record is contractive force; bottom record is simultaneous extracellular recording from posterolateral connective. C2: peak response to perfusion of 1 µM DA in same preparation as C1. Note that the intensity of motor neuron activity is not enhanced. (D) Effect of DA to accelerate and enhance motor neuron activity in the isolated CG preparation. D1: top record is intracellular recording from CG motor neuron; bottom record is simultaneous extracellular recording from posterolateral connective. D2: peak response to perfusion of 1 µM DA in the same preparation as D1. (E) Model of DA release from L cell terminals into the pericardial organ, where it reaches heart muscle indirectly as a hormone (dotted arrow with question mark) and by direct innervation into the CG (solid arrows). The innervation consists of a posterior projection to the posterior pacemaker interneurons (thin solid arrow) and an anterior projection to the motor neurons (thick solid arrow). A peripheral feedback mechanism blocks DA enhancement of CG motor neuron burst duration. B–E from Fort et al. 2004, with permission from The American Physiological Society.

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Ronald Harris-Warrick A large number of peptides have been identified by their effects on the crustacean heart (Christie et al. 2010a); as with octopamine, their effects may vary between species. The first peptide studied, proctolin, is a powerful cardioaccelerator in H. americanus, when applied to the whole CG or selectively to either the pacemaker small cells or the large motor neurons (Sullivan and Miller 1984). Proctolin evokes driver potentials in silent, tetrodotoxin (TTX)-treated large cells. This large inward current has been argued to arise from a conductance decrease to potassium (Sullivan and Miller 1984) or activation of a voltage-dependent sodium current (Freschi 1989). A  number of other peptides released from the PO and elsewhere affect heart rate (reviewed by Christie et  al. 2010a). In the crab C.  borealis, of 10 peptides tested, eight increased burst frequency and motor neuron spike frequency in the isolated CG, including red pigment-concentrating hormone (RPCH), tachykinin-related peptide (CabTRP Ia), and a series of extended FLRFamide peptides. In contrast, only one tested peptide, allatostatin III type A, inhibited CG motor output (Cruz-Bermúdez and Marder 2007). Saver et al. (1999) studied the effects of several peptides on neurons isolated from the Carcinus maenas CG: crustacean cardioactive peptide (CCAP), the FLRFamide-related peptide F2, and proctolin all initiated or accelerated rhythmic bursting by an increase in the slope of the pacemaker potential that initiates the bursts. Many other peptides have cardioactive effects, but little is known of their mechanisms of action. A number of these cardiac neuromodulators also have direct effects on the cardiac muscle, typically altering the strength of nerve-evoked contractions, and this can indirectly affect cardiac cycle frequency via modulation of sensory feedback from the muscle to the CG. Wilkens et al. (2005) showed that proctolin and the FLRF-related peptide F2 both directly enhance the amplitude of electrically evoked contractions, and at higher concentrations cause a tonic contraction independently of nerve stimulation. Calcium imaging studies suggest that these peptides enhance voltage-dependent Ca 2+-induced calcium release from the sarcoplasmic reticulum to enhance electrically evoked contractions, while directly gating sarcolemma Ca 2+ current at higher concentrations to evoke tonic contractions. In the embryonic and juvenile stages of the isopod Ligia exotica, the heart rhythm is myogenic; in early juveniles, DA decreases the frequency and increases the action potential duration, slowing the beat frequency, while increasing the force of the contractions (Yamagishi et al. 2004); the DA effect reverses to a positive chronotropic effect in older juveniles. The actions described above are examples of extrinsic neuromodulation by amines and peptides that are not themselves expressed by CG neurons. Cropper et al. (1987) and Katz (1995) introduced the distinction between extrinsic modulatory inputs that are optional to modify network output, and intrinsic modulatory inputs that are released by network neurons and thus obligatorily activated whenever the network is active, helping to shape its output and its modification over time. There has been considerable discussion over whether there is intrinsic neuromodulation in the CG system. Recent molecular analysis shows that the gene for the peptide calcitonin-like diuretic hormone is expressed as RNA in CG motor neuron somata in H. americanus (Christie et  al. 2010b). This peptide increases heart contraction frequency and amplitude at very low concentrations, and may be the first intrinsic peptide in the CG. Acetylcholine (ACh) is not used as a neurotransmitter in the CG or on the heart muscle, yet the CG can synthesize ACh and contains acetylcholinesterase (Sullivan and Miller 1990); when bath-applied, it increases the strength and speed of heart contractions by activating a complex net inward current (Freschi and Livengood 1989). Finally, modulatory actions in the crustacean heart may result from a complex feedback interaction between actions at the CG and at the heart muscle. For example, nitric oxide (NO) appears to be a retrograde transmitter, synthesized and released from heart muscle during ongoing calcium-evoked contractions, and having an effect on the CG to slow and weaken heart



Modulation of Crustacean Networks for Behavior

contractions (Goy 2005). The weakened strength appears to be an indirect consequence of loss of synaptic facilitation at the neuromuscular junction at the slower cycle frequency. In the CG, NO does not appear to act via activation of soluble guanylate cyclase; indeed, cyclic guanosine monophosphate analogs increase rather than decrease the cycle frequency and contraction amplitude (Goy 2005). Miller and colleagues have studied the interaction of central and peripheral modulatory effects by independently applying DA (Fort et al. 2004) as well as several peptides to the isolated CG or to the semiintact or intact heart in the crab Callinectes sapidus. These compounds have direct actions both on the CG and on the muscles to exert positive chronotropic and inotropic actions. In the case of DA, a pair of commissural ganglion (CoG) dopaminergic neurons, the L cells, send an axon that terminates around large and small cells in the CG (Fig. 5.1B). There are no direct synapses onto the cardiac muscle. However, the L-cell also sends major processes to the PO, where it releases DA into the hemolymph to act as a neurohormone on both the CG and the heart muscle (Fig. 5.1E). Comparison of the effects of DA on the intact or semi-intact heart versus the isolated CG indicates both direct effects on the CG to increase the cycle frequency and direct effects at low, hormonal concentrations to enhance nerve-evoked muscle contraction (Fig. 5.1C, D). Dopamine also modulates neuronal burst properties in the isolated CG that are suppressed by sensory feedback in the intact system, which appears to arise from stretch-sensitive terminals of the motor neurons themselves (Fig. 5.1E). This work enlarges the concept of appropriate modulation of the neuromuscular transform (Brezina et al. 2000), where the properties of the central motor commands and the nonlinear response properties of the peripheral musculature must be correctly matched to drive appropriate behavior. Here, coordinated modulation by the dual release of DA within the CG and from the PO to the heart muscle assures appropriate cardioacceleration with the correct muscle properties.

MODULATION OF THE SWIMMERET SYSTEM The swimmerets (also called pleopods) are a set of paired appendages on the underside of each segment of the crustacean abdomen. They move rhythmically in a reverse metachronous wave during forward swimming and certain other behaviors, driven by sets of power stroke (PS) and return stroke (RS) motor neurons located in the abdominal ganglia. A distributed CPG in the abdominal ganglia, which includes a set of nonspiking interneurons, organizes the alternating pattern of PS and RS motor neuron bursts, and intersegmental coordinating neurons set up the delay between segments (Mulloney and Smarandache-Wellmann 2012). Studies of neuromodulation in this system have focused primarily on identifying compounds that can initiate or terminate the swimmeret motor pattern in the isolated nerve cord of the crayfish Pacifastacus leniusculus. Early work showed that the cholinergic agonist carbachol can initiate the swimmeret rhythm, acting primarily via modulatory muscarinic ACh receptors but with a nicotinic enhancement of burst frequency in active preparations (Braun and Mulloney 1993). The peptides proctolin and CCAP can also activate a silent swimmeret network (Mulloney et al. 1997), but they activate different subsets of motor neurons to generate somewhat different patterns. In contrast, octopamine inhibits ongoing swimmeret activity (Mulloney et  al. 1987). Recent voltage clamp analysis shows that both carbachol and octopamine directly affect swimmeret motor neurons (Tschuluun et  al. 2009). Carbachol evokes a slow inward current to activate the swimmeret motor pattern. A portion of this current is blocked in low calcium/ high magnesium saline, but it is unclear whether this results from block of a carbachol-activated calcium or calcium-dependent inward current, or synaptic block of a tonic depolarizing current from unidentified premotor interneurons. Octopamine inhibits the neurons by activating an

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Ronald Harris-Warrick outward current mediated by a conductance increase. These currents have not been further characterized.

MODULATION IN THE STOMATOGASTRIC NERVOUS SYSTEM The crustacean STG has long served as a major model for studying the inner workings of neural networks for behavior. This tiny ganglion, which has between 26 and 30 neurons in different species, contains two complete CPG networks (the gastric mill and the pyloric CPGs), and interacts with several CPGs located in other ganglia. Electrophysiological studies of the STG began with the groundbreaking work of Donald Maynard and was brought to the main stage by Allen Selverston, whose insightful review in 1976 (Selverston et al. 1976), introduced the STG to the general neuroscience audience. For over 40 years, the pyloric and gastric mill CPGs have been the best understood and most thoroughly studied networks in neuroscience. Stein (2009) has written an excellent and comprehensive review of modulation in the stomatogastric nervous system (STNS). Chapter 13 by Marder gives a detailed description of the stomatogastric system, so only a brief introduction to the pyloric and gastric mill CPGs is given here. The pyloric CPG drives rhythmic pumping and filtering movements of the posterior, or pyloric, region of the crustacean foregut. It contains between 11 and 14 neurons in different species, of which one is an interneuron while the remaining are both constituents of the CPG network and motor neurons that drive muscle contractions. The triphasic rhythm is driven primarily by the conditional pacemaker anterior burster (AB) interneuron, electrically coupled to two pyloric dilator (PD) motor neurons; this pacemaker kernel bursts rhythmically at 0.5–2 Hz, and inhibits all the other neurons. Following the burst, the follower neurons recover to resume firing at different phases due to intrinsic differences in their rates of postinhibitory rebound and the pattern of synaptic inhibition within the network. The slower gastric mill CPG drives the movement of the lateral and medial teeth inside the foregut in several different phasing rhythms, depending on the modulatory milieu. It contains between nine and 11 neurons, including one interneuron. None of the neurons plays a unique pacemaker role. Instead, several pairs of mutually inhibitory neurons coordinate a basic biphasic pattern, with a frequency about one tenth of the pyloric rhythm (for further details, see c­ hapter 13 in this volume). Sources of Modulatory Input to the Stomatogastric Ganglion There are three major sources of modulatory inputs to the STG (see Fig. 13.3 in this volume). First, the STG is located within the anterior ophthalmic artery, so circulating neurohormones can modify network function at the low (typically nM or lower) concentrations seen in the hemolymph. At least 28 compounds affect the STG as neurohormones, including 5HT, octopamine, DA, and at least 25 neuropeptides (Marder and Bucher 2007, Christie et al. 2010a). These are released from several known neurosecretory organs including the adjacent PO, the postcommissural organ, the X-organ/sinus gland complex in the eyestalk, and the anterior cardiac plexus (Christie et al. 2010a). Second, the STG is connected to the rest of the nervous system by a single nerve, the stomatogastric nerve (stn), which contains the axons of projection neurons providing modulatory input to STG networks. Although it carries the thin ascending axons of many sensory neurons, the stn contains relatively few descending axons:  in the California spiny lobster P.  interruptus the stn contains about 110 large descending axons (King 1976), while in the crab C. borealis there are only about 40 descending axons (Coleman et  al. 1992, M.  Nusbaum personal



Modulation of Crustacean Networks for Behavior

communication). Nusbaum and coworkers have identified a number of projection neurons in the crab C. borealis: four pairs of identified neurons (MCN1, MCN5, MCN7, and CPN2) are found in the CoG, while the proctolin-containing modulatory proctolinergic neuron (MPN) is found in the oesophageal ganglion (OG); the IVN neurons are found in the brain (Stein 2009). Homologs of these neurons are found in other crustacean species, along with some novel neurons (Stein 2009). Many additional projection neurons wait to be identified. These neurons each release multiple neurotransmitters, typically combining small transmitters with peptides, and exert a variety of different actions to shape STG network function (Nusbaum and Beenhakker 2002). Third, the STG and the projection neurons in higher ganglia receive input from peripheral modulatory/sensory neurons. These neurons are typically mechanoreceptors monitoring muscle stretch or tendon organ tension. The first identified modulatory/sensory neurons were the GPR neurons, which in C. borealis contain ACh, 5HT, and the peptide allatostatin (Katz et al. 1989, Skiebe and Schneider 1994). These neurons have a complex set of effects both directly on CPG neurons in the STG (Katz and Harris-Warrick 1990) and indirectly, by activating modulatory projection neurons in higher ganglia (Blitz et al. 2004). They may also modulate their own sensitivity to stretch (Birmingham et  al. 2003). The anterior gastric receptor (AGR) neuron appears to have no direct synapses in the STG but synapses on higher projection neurons and can activate or entrain the gastric mill rhythm (Smarandache and Stein 2007). Several major points can be made concerning these varying sources of modulatory input to the STG. First, modulators can affect CPG activity both by direct actions onto the CPG network neurons in the STG, and indirectly by affecting the modulatory projection neurons. As mentioned above, the GPR neurons act both directly on CPG neurons in the STG and indirectly on projection neurons in the CoG, while the AGR neurons act only indirectly in the CoG. More than 75 neurons project to the CoG from the brain, and it is likely, though not yet proven, that many of these modulate the projection neurons to modify STG network activity (Stein 2009). The IVN neurons in C. borealis and P. interruptus act at different locations to inhibit the pyloric rhythm and excite the gastric mill rhythm (Hedrich and Stein 2008). Inhibition of the pyloric network results from IVN release of histamine within the STG (Claiborne and Selverston 1984), while gastric mill excitation results from IVN release of an FLRFamide-related peptide in the CoG, which elicits bursting in some modulatory projection neurons (Hedrich and Stein 2008, Stein 2009). The actions of many neurotransmitters have only been studied in the isolated STG (Marder and Bucher 2007); it is not known how many of these transmitters simultaneously affect projection neurons in the CoG and elsewhere. Second, the same compound can act both as a neurohormone, reaching its targets via the hemolymph, and as a transmitter released from modulatory neurons. For example, in the crab, DA, 5HT, octopamine, and at least 16 peptides are released both into the hemolymph from neurosecretory sites to affect the STG, and from nerve terminals of either projection neurons or sensory/modulatory neurons within the STG (Marder and Bucher 2007). It is likely that these paracrine and neurohormonal actions are coordinated, as described above for the L cell and its combined hormonal and transmitter actions in the CG. Third, circulating neurohormones may modify the activity of modulatory projection or sensory neurons, altering their effects on STG networks. Most research on neuromodulator action has used a single modulator or projection neuron at a time, but recent studies have started to look at their complex interactions when simultaneously active. For example, DeLong and Nusbaum (2010) showed that stimulation of the sensory/modulatory GPR neurons alone prolongs the retractor phase of the gastric mill rhythm via a presynaptic inhibition of the projection neuron, Modulatory Proctolin Neuron MCN1. If the peptide neurohormone CCAP is applied, it reduces the effect of GPR by a complex effect on inputs to MCN1 that negates the impact of

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Ronald Harris-Warrick the GPR neurons. Thus, the effect of GPR on the gastric mill motor pattern is regulated by a circulating neuropeptide. Fourth, there are species differences in which compounds are hormones or transmitters in the STNS. For example, in the crab C. borealis, the GPR neurons are the sole neuronal source of 5HT to the STG, where it acts at micromolar concentrations on the STG neurons (Katz et al. 1989). In contrast, the GPR neurons in the spiny lobster P. interruptus lack 5HT, and there is no neuronal 5HT input to the STG; in the lobster, 5HT acts only as a neurohormone at nM concentrations (Beltz et al. 1984). Fifth, there is growing evidence for both intrinsic and extrinsic modulatory actions in the STG. Acetylcholine and glutamate are the neurotransmitters of all STG neurons, and act on ionotropic receptors to shape the motor pattern (Harris-Warrick et al. 1992). However, in four species of crayfish, one or more neurons show immunolabeling for an allatostatin-like peptide (Skiebe 1999). This is not seen in other species, and apparently has not been pursued. In the shrimp Palaemon serratus, several FMRFamide-immunoreactive neurons are seen in the STG; they send axons to the PD muscles, though they are not the PD motor neurons (Meyrand and Marder 1991); these neurons are not found in other species. With regard to modulatory effects of glutamate and ACh, Nagy’s group (Nagy and Dickinson 1983) described the muscarinic actions that excite the pyloric network in Jasus lalandii, and Selverston and colleagues have provided excellent evidence for metabotropic glutamatergic receptor actions in P. interruptus, which can affect the gastric mill network (Krenz et al. 2000). However, there is as yet no direct evidence that glutamate or ACh, released directly from the STG neurons, has access to these modulatory receptors (Katz and Harris-Warrick unpublished, A. Selverston personal communication). Instead, extrinsic modulatory projection neurons may normally activate them. Different Responses by Application of Transmitters versus Stimulation of Modulatory Neurons These two methods of delivering modulatory inputs to the STG networks have provided interesting and complementary data. Most mechanistic studies have used bath application or iontophoresis of the modulator. This allows the researcher to study the effects of one compound at a time, at known concentrations, or with known time course of delivery. This approach is optimal for understanding the cellular mechanisms of modulator action, and to identify the neuronal, ionic, and molecular targets of neuromodulators in a network. Bath application also mimics the actions of circulating neurohormones. Stimulation of modulatory neurons is the only way to mimic both the complement of transmitters coreleased by the neuron and the spatial and temporal properties of the released transmitters. The exact temporal pattern of stimulation may matter in some cases but not others. For example, the actions of released 5HT from the sensory/modulatory GPR neurons in the STG are mimicked by bath application of 5HT (Katz and Harris-Warrick 1989), but GPR presynaptic inhibition of the projection neuron MCN1 depends on the phase; GPR activation during the retraction phase of the gastric mill rhythm delays the switch to protraction (DeLong et al. 2009). Wood et al. (2004) showed that tonic and rhythmic MCN1 stimulation evoke somewhat different gastric mill patterns. Tonic stimulation activates the gastric mill rhythm. However, in the intact system, the MCN1 neuron receives periodic pyloric feedback that causes it to fire in a pyloric pattern; this rhythmic MCN1 firing pattern evokes a gastric mill rhythm with much more marked pyloric-timed interruptions of some of the gastric mill neuron bursts. The net effect of bath-applying a transmitter may be qualitatively different from the same transmitter when released from modulatory neuron terminals. In part, this reflects the multiplicity of cotransmitters that are coreleased from the terminal. However, more sophisticated



Modulation of Crustacean Networks for Behavior

differences are seen. For example, there are marked differences in the strength of the proctolin effect when released from the MPN and the MCN1 projection neurons onto their STG targets, which is not seen when proctolin is bath-applied (Wood and Nusbaum 2002). This arises from the presence of extracellular peptidases that degrade proctolin after release; peptidase blockade reduces the differences between the two neurons, and between the neurons and bath-applied proctolin. Thus, extracellular peptidases sculpt the actions of the neurally released neuropeptide, presumably by limiting its range of diffusion from its release sites to distant receptors (Wood and Nusbaum 2002). Oginsky et al. (2010) found that the distribution of D2α DA receptors is not uniform over the PD neuron: they are concentrated in the synaptic neuropil, but are only present at 40% of the defined synaptic structures of any PD neuron. Synaptically released DA may not access all of these structures. The effects of modulatory compounds or neurons can be studied at the network, cell, and synaptic levels of analysis. Below, I discuss modulation in the STG at each of these levels. Modulation at the Network Level One of the major principles to emerge from studies of modulation in the STNS is that an anatomically defined neural network can generate a variety of related motor patterns, due to the actions of neuromodulators (Fig. 5.2). Both the pyloric network and the gastric mill network show great flexibility in the motor patterns they produce when different modulatory inputs are active. Neuromodulators generate this flexibility by several linked mechanisms: 1. Modulatory input can initiate the motor pattern from a silent state or terminate the pattern from an active state. 2. Modulatory inputs can modify the cycle frequency, via both direct effects on the major pacemaker kernel (Ayali and Harris-Warrick 1999) and indirect effects on neurons that provide feedback to the pacemaker kernel (Ayali and Harris-Warrick 1999, Johnson et al. 2011). 3. Modulatory inputs can alter the phasing of the motor pattern, by differential actions on the intrinsic firing properties of the neurons and the strengths of their synaptic interaction. 4. Modulatory inputs can alter the activity of each network neuron, ranging from strong activation to complete inhibition, as well as altering its baseline intrinsic properties to enable bistability, pacemaker activity, and variable postinhibitory rebound. 5. In this way, neuromodulators can select which neurons are actively participating in the network at any time: by differential effects on CPG neurons, they can define the neuronal composition of the functional circuit from the anatomically defined network. Several additional principles have been demonstrated from studies of STG network modulation. First, neuromodulators can have either convergent or divergent actions on the neurons in a network. For example, muscarinic agonists and a number of peptides (including proctolin, CCAP, and FLRFamide-related peptides) appear to converge onto activation of a single inward excitatory current, called I MIC, that excites neurons within the physiological voltage range (Swensen and Marder 2000) (Fig. 5.2B). However, the motor patterns generated by these compounds are very different. This arises primarily from the differential expression of peptide receptors on different network neurons (Swensen and Marder 2000) (Fig. 5.2C). By activating different subsets of network neurons, each peptide can generate a different motor pattern. In an interesting proof of this concept, Swensen and Marder (2001) compared the pyloric motor patterns elicited by CCAP and proctolin. Both peptides activate I MIC, but proctolin excites three neurons that are not responsive to CCAP. Swensen and Marder used the dynamic clamp

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Fig. 5.2. Multiple pyloric motor patterns evoked by peptides and muscarinic agonists by convergence on a single ionic current. (A) Simultaneous recordings from the lateral pyloric (LP) and pyloric dilator (PD) motor neurons with nerve recording from the lateral ventricular nerve (lvn), which carries axons of the LP, PD, and pyloric (PY) neurons. Each modulator was added to a silent preparation and evoked a unique motor pattern (from Marder and Thirumalai 2002, with permission from Elsevier Press). (B) Peptides and muscarinic agonists activate receptors that converge to activate a single ionic current, I MIC (from Marder and Bucher 2007, with permission from The American Physiological Society). (C)  Each peptide activates I MIC in a different subset of pyloric neurons, thus generating different motor patterns (from Swensen and Marder 2001, with permission from The Society for Neuroscience). Neuron names: AB, anterior burster; IC, inferior cardiac; LP, lateral pyloric, PD, pyloric dilator; PY, pyloric constrictor; VD, ventricular dilator.



Modulation of Crustacean Networks for Behavior

to artificially add I MIC to two of these three neurons in the presence of CCAP, and effectively converted the CCAP-evoked motor pattern to the proctolin pattern. In contrast, detailed studies of the cellular mechanisms of pyloric network modulation by the monoamines DA, 5HT, and octopamine show significant divergence in mechanisms of action (Harris-Warrick et  al. 1998, Harris-Warrick and Johnson 2010)  (Fig.  5.3). Each of these amines affects all or nearly all of the neurons in the pyloric network (Flamm and Harris-Warrick 1986). However, the effects of each amine are different on different neurons. For example, DA induces rhythmic oscillations in the AB neuron, inhibits and hyperpolarizes the PD and ventricular dilator (VD) neurons, and excites tonic activity in the lateral pyloric (LP), pyloric (PY), and inferior cardiac (IC) neurons (Fig. 5.3B). In contrast, octopamine excites all of the neurons, but with different strengths and effects: it induces bursting in the AB neuron and strongly excites the LP and PD neurons, so that the other neurons are synaptically inhibited and only fire weakly in the motor pattern. Serotonin evokes rhythmic bursting in the AB neuron and tonic activity in the IC neuron, but inhibits the LP and VD neurons, with no apparent effects on the PD and PY neurons. As a consequence, each amine evokes a motor pattern with different neurons active or silent, sculpting out different functional circuits from the anatomically defined network. These divergent effects are mediated by the multiplicity of receptors that the monoamines can activate. Zhang and Harris-Warrick (1994) used pharmacological tools to show that the burst-inducing, excitatory, and inhibitory actions of 5HT are mediated by different receptors. Recently, Baro and colleagues showed that neuron-specific expression of D1 and D2 DA receptors can explain the different neuronal responses to DA. DA excites the LP neuron, which only expresses D1 receptors that activate the cAMP pathway; in contrast, DA inhibits the PD neuron, which only expresses the D2 receptor that suppresses the cAMP pathway (Oginsky et  al. 2010, Zhang et al. 2010). The different receptors can activate different patterns of ionic currents that shape the neuronal activity (see below). Second, modulatory projection neurons can evoke unique motor patterns from the STG networks (Fig. 5.4). In part, this is because they express different (though partially overlapping) sets of cotransmitters (Nusbaum and Beenhakker 2002, Stein 2009). For example, the MPN neurons use proctolin and gamma-aminobutyric acid (GABA) as cotransmitters. The MCN1 neurons also contain proctolin and GABA, but in addition are the only neurons to use cancer tachykinin-related peptide (CabTRP) Ia as a transmitter. MCN7 also uses proctolin (Stein 2009). Each of these neurons evokes different effects on the gastric mill and pyloric rhythms (Wood et al. 2000) (Fig. 5.4A). Even after nonproctolin actions are pharmacologically blocked, these neurons still evoke different motor patterns, in part due to spatial sculpting by extracellular peptidases described above (Wood and Nusbaum 2002). Third, even though a modulatory neuron releases a set of cotransmitters and STG neurons express receptors for those transmitters, not all the neurons respond to the released transmitters. For example, MCN1 coreleases CabTRP Ia, proctolin, and GABA. The gastric mill Dorsal Gastric (DG) neuron responds to both peptides when bath-applied, but MCN1-evoked DG excitation is mediated exclusively by CabTRP Ia (Stein et al. 2007). The Lateral Gastric (LG) neuron expresses receptors for both CabTRP Ia and GABA, but MCN1’s effects on LG are also exclusively mediated by CabTRP Ia. MCN1 excites Int1, but this excitation is not mediated by either peptide, and is exclusively mediated by GABA (Stein et al. 2007). In the crab, the sensory-modulatory GPR neurons contain ACh, 5HT, and allatostatin, but its effects on the pyloric network in the STG are mediated only by ACh and 5HT, with no detectable effect of allatostatin (Katz and Harris-Warrick 1989). It is not clear how these selective responses to a subset of a neuron’s transmitters is achieved, though a spatial separation of the receptors from the release sites is an obvious possibility (Stein 2009).

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Ronald Harris-Warrick Fourth, several studies have shown that the effects of a modulator on a network are state dependent: both excitatory and inhibitory modulatory effects are typically stronger if the starting motor pattern is weak (Christie et al. 2010a). Indeed, some neuromodulators have very little effect if the motor pattern is strong. For example, proctolin can increase the pyloric cycle frequency, but only up to around 1 Hz; if the preparation is cycling more rapidly, proctolin does not further increase the frequency (Hooper and Marder 1987). The inhibitory effects of allatostatin are also more marked on a slowly cycling preparation than a rapidly cycling one (Skiebe and Schneider 1994). The mechanism underlying this state dependence has not been elucidated, but presumably reflects the simultaneous actions of other neuromodulators on the system, whose interactions may not be strictly summative. For example, another motor network drives slow rhythmic contractions of the cardiac sac. Dickinson et al. (1997) showed that the peptide RPCH can activate the cardiac sac rhythm in the isolated STG-OG preparation, but proctolin cannot. However, shortly after termination of an RPCH-evoked cardiac sac episode, proctolin can activate a similar motor pattern, and subthreshold concentrations of the two peptides together can

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Fig. 5.3. Divergent actions of monoamines on the pyloric network. (A) Dopamine, serotonin, and octopamine each evoke a different variant of the ongoing pyloric motor pattern (modified from Ayali and Harris-Warrick 1999, with permission from The Society for Neuroscience). (B) Different effects of DA in each pyloric neuron when isolated from all synaptic input. Dopamine evokes rhythmic bursting, tonic activity, or inhibition in different neurons. Voltage clamp studies show that DA affects different ionic currents, in different directions, in each neuron (figure modified from Flamm and Harris-Warrick 1986, with permission from The American Physiological Society). Neuron names: AB, anterior burster; IC, inferior cardiac; LP, lateral pyloric, PD, pyloric dilator; PY, pyloric constrictor; VD, ventricular dilator.



Modulation of Crustacean Networks for Behavior

activate the cardiac sac rhythm. This effect appears to be mediated by strong enhancement of the IVN excitatory post-synaptic potentials (EPSPs) within the STG. Finally, modulatory inputs can completely reconfigure several CPG networks, causing them to fuse into novel conjoint networks (Dickinson and Moulins 1992, Dickinson 1995) (Fig. 5.4). Stimulation of the proctolinergic MCN1, MPN, or MCN7 neurons not only differentially affects the pyloric network, as described above, but also alters interactions between the pyloric and gastric mill networks: MCN1 and MCN7 activate different patterns of the gastric mill network, and these have different effects on timing of pyloric neuron activity (Fig. 5.4A). Neurons that are components of one network under one condition may switch to fire with another network under other conditions. For example, in Palinurus vulgaris, the VD neuron is active with the pyloric pattern, but when the cardiac sac pattern becomes active, VD switches and fires in cardiac sac time, due to strong synaptic drive from the IVN neurons (Hooper and Moulins 1990). Weimann et al. (1991) found that different neurons had varying propensity to switch between the gastric mill and pyloric networks in the crab, with some never switching while others frequently switch; they propose that the neurons should be combined into a “gastropyloric pool” from which different modulatory inputs can select components for the gastric or pyloric networks. The gaseous transmitter NO may help to maintain the separation between the gastric mill and pyloric networks (Scholz et al. 2001): in the crab Cancer productus, when NO activity is blocked, the gastric mill rhythm is disrupted and several of its neurons join the pyloric network. More dramatic network fusion can occur when modulatory inputs are activated. For example, RPCH fuses the gastric mill and cardiac sac networks to generate a motor pattern that is completely different from either parent network (Dickinson et al. 1990) (Fig. 5.4B). The most dramatic fusion occurs after stimulation of the pyloric suppressor neurons in Homarus gammarus (homologs of the IVN neurons in other species), when the pyloric, gastric mill, and oesophageal networks are all disrupted and fuse to form a conjoint motor pattern that has been interpreted as a swallowing rhythm (Meyrand et al. 1991) (Fig. 5.4C,D). This fusion occurs via a complex set of modulatory effects with different kinetics on the intrinsic firing properties and synaptic interactions between the networks (Faumont et al. 2005). Modulation at the Level of the Neuron Each neuron displays a set of intrinsic electrophysiological properties that determine its baseline activity at any moment; this is determined by the set of ionic currents that are active at the moment. Neuromodulators alter the strength and properties of these currents, allowing the CPG neurons to display many different firing patterns. Modulation of Bursting Oscillations All the pyloric neurons are conditional oscillators and can generate rhythmic bursts of action potentials only when appropriate modulatory inputs are present. When modulatory input to the STG is completely blocked, the pyloric network slows down and, in P. interruptus, stops altogether (Russell and Hartline 1978). As the most rapid oscillator, the AB neuron is the major pyloric pacemaker. However, the AB can oscillate at a variety of frequencies. Dopamine, 5HT, and octopamine all elicit bursting with different frequencies and amplitudes in the silent, synaptically isolated AB neuron (Flamm and Harris-Warrick 1986, Harris-Warrick and Flamm 1987) (Fig. 5.3B). Pharmacological and voltage-clamp experiments show that this neuron uses very different ionic mechanisms to induce bursting with each monoamine. DA-evoked bursting is dependent on calcium but not on sodium currents; it evokes the release of calcium from intracellular stores, which is essential for DA-evoked bursting to occur (Kadiri et al. 2011). This

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Fig. 5.4. Modulation of interactions between nominally separate neural networks. (A) Motor patterns evoked by stimulation of MCN1, MCN7, and MPN projection neurons. All release proctolin but differ in additional transmitters. They also differ in the neurons affected by each transmitter and thus evoke motor patterns with varying interactions between the pyloric and gastric mill networks (from Nusbaum and Beenhakker 2002, with permission from The Nature Publishing Group). (B) Application of red pigment concentrating hormone (RPCH) causes a fusion of the cardiac sac and gastric mill networks. Top panel: Recordings from a gastric mill neuron, LPG, and a cardiac sac neuron, CD2, under control conditions showing a large difference in cycle frequency. Middle panel: After addition of RPCH, the networks fuse to a new conjoint network with an intermediate frequency. Bottom panel: Schematic showing the fusion of the two networks (from Dickinson et al. 1990, with permission from The Nature Publishing Group). (C) Stimulation of the pyloric suppressor (PS) neuron evokes a conjoint network with components from the oesophageal, gastric mill and pyloric networks. Recordings from gastric mill neurons (MG and LG) and pyloric neurons (PY and PD) under control conditions showing very different motor patterns and cycle frequencies. During and following PS stimulation, these neurons fire in a unique joint pattern with intermediate cycle frequency (from Faumont et al. 2005, with permission from John Wiley and Sons). (D) Model of the fusion of the three networks into the conjoint swallowing network, and its further change with time (from Meyrand et al. 1994, with permission from The Society for Neuroscience).

in turn activates a calcium-activated nonselective current, ICAN, which provides the depolarizing ramp to begin each burst. The accumulation of calcium activates a calcium-activated potassium current, I K(Ca), which is thought to terminate the burst and hyperpolarize the neuron (Gola and Selverston 1981, Kadiri et al. 2011). Dopamine alters a number of other currents that support AB bursting, including a decrease in the transient potassium current, I A , and a leak current, and an increase in the hyperpolarization-activated inward current, I h (Harris-Warrick and Johnson 2010). Paradoxically, DA also reduces the voltage-activated calcium current, ICa(V), which normally assists in bursting in other neurons, and enhances a slowly activating delayed rectifier potassium current, I K(V), which would normally slow the spike frequency during the



Modulation of Crustacean Networks for Behavior

burst (Harris-Warrick and Johnson 2010) (Fig. 5.3B). In contrast, 5HT-evoked bursting is not mediated by release of calcium from intracellular stores. Instead, it is abolished by TTX and low sodium, and relies on a persistent sodium current to generate the ramp depolarization to initiate the burst (Harris-Warrick and Flamm 1987, L. Kadiri and R. Harris-Warrick unpublished). Modulation of Plateau Potentials Many of the gastric mill and some pyloric neurons show bistable properties in the presence of the appropriate neuromodulators (Hartline and Graubard 1992). In this state, a brief depolarization evokes a prolonged depolarized “plateau potential” during which the cell fires tonic action potentials. Bistability converts the role of synaptic input from that of a driver, which determines the strength of the postsynaptic response, to a simple trigger that initiates the intrinsic prolonged response. Bistability is conditional in STG neurons: it is lost when modulatory inputs are removed. The mechanisms of plateau induction are poorly understood in most cases. Stimulation of the cholinergic/serotonergic GPR neurons can evoke a plateau potential in the crab DG neuron by a dual mechanism of rapid nicotinic depolarization to initiate firing and a slow serotonergic depolarization that maintains the plateau (Kiehn and Harris-Warrick 1992a, 1992b). Serotonin’s mechanisms for inducing the bistability are complex. The critical step is an enhancement of ICa(V) that in turn triggers ICAN. Activating ICAN alone by light-induced release of intracellular caged calcium is sufficient to trigger the full plateau potential (Zhang and Harris-Warrick 1995, Zhang et al. 1995). Serotonin also modifies other currents that support the plateau, including enhancement of the hyperpolarization-induced inward current, I h, and reduction of I K(Ca) (Kiehn and Harris-Warrick 1992a). Other mechanisms support bistability in different neurons. For example, Elson and Selverston (1997) presented evidence that a persistent sodium current, I NaP, underlies plateau potentials in the LG and LPG neurons in P. interruptus. Modulation of Postinhibitory Rebound and Delayed Excitation Most of the pyloric neurons show a complex response to a hyperpolarizing step: a delayed repolarization is followed by a slower depolarizing overshoot that evokes spiking. This mechanism of postinhibitory rebound (PIR) helps the neurons to recover from synaptic inhibition and initiate the next burst cycle (Hartline and Graubard 1992). The ability of synaptic inhibition to evoke a delayed excitation uncovers the dual nature of inhibition in many neural networks. The amplitude and rate of PIR helps determine the phasing of neuronal activity in the motor cycle, which is essential in shaping the motor pattern. The ionic mechanisms underlying PIR and delayed excitation are complex. Two major currents shape the recovery from inhibition in pyloric neurons. The I A is inactivated at depolarized voltages, and de-inactivated by hyperpolarization, allowing it to be transiently activated following the hyperpolarization, slowing the rate of PIR. The I h is slowly activated during hyperpolarization providing a slowly deactivating depolarizing driver current that sustains the depolarizing overshoot driving PIR (Harris-Warrick et al. 1995b). In other systems, a low-threshold T-type calcium current also contributes to PIR. Postinhibitory rebound is subject to neuromodulation, which can change the phasing of neuronal bursts in the pyloric rhythm. Harris-Warrick et al. (1995a, 1995b) analyzed the mechanisms by which DA causes phase advances in the LP and PY neurons (Fig. 5.3B). Dopamine accelerates the rate of PIR in a subset of the PY neurons by directly reducing I A in several ways. It selectively reduces the conductance of a rapidly inactivating component of I A , and accelerates the rate of inactivation; DA also shifts the voltage activation and inactivation curves to the right, reducing activation with subthreshold depolarizations. There is little or no contribution of I h to PIR in PY neurons (Harris-Warrick et al. 1995a). The LP neuron is also phase-advanced by DA, but two

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Ronald Harris-Warrick different mechanisms contribute to this phase change (Harris-Warrick et al. 1995b). First, as in the PY neurons, DA decreases I A , but with subtle differences in the biophysical mechanism. In the LP neuron, DA reduces the maximal conductance of both rapidly and slowly inactivating components of I A , in addition to depolarizing the voltage dependence of activation and inactivation to a greater extent than in the PY neurons. Second, DA enhances I h by shifting its voltage activation curve to a more depolarized range, an effect which does not occur in PY neurons. Thus, a complex and variable opposition between I h and I A helps to determine the onset phasing of the LP neuron. Modulation of Spike Frequency and Timing In STG networks, most of the neurons are simultaneously components of the CPG and motor neurons that carry the network output to the muscles. Thus, the pattern and frequency of action potential firing will affect muscle contraction and the resulting behavior. Most modulatory inputs act to increase or decrease spike frequency in their target neurons. The ionic mechanisms underlying the regulation of spike frequency are not well understood. For the pyloric LP and PY neurons, DA enhances spike frequency in part by reducing I A (Harris-Warrick et al. 1995a, 1995b) (Fig. 5.3B). Muscarinic agonists and a number of peptides converge on a single excitatory current, I MIC . This inward current, carried predominantly by Na+, shows a U-shaped I/V relation, with maximal activation around -40 mV, due to calcium-dependent rectification at hyperpolarized voltages (Golowasch and Marder 1992). This makes it maximally effective to depolarize the neuron near the threshold for spike generation (Stein 2009). The ion channels for this unusual current have not been identified, but Zhao et al. (2010) used modeling to suggest that it may arise from changes in leak currents, including a conductance decrease of a leak current with a reversal potential slightly below the resting potential range. Regulation of spiking occurs at unexpected locations in the stomatogastric system. Several neurons have multiple spike initiation zones (SIZs), one of which is in the STG while another is along the axon at a significant distance, and is selectively activated by modulatory input. For example, in H. americanus, the peripheral axons of the pyloric PD neuron are excited by nanomolar concentrations of DA, resulting in tonic spike activity not related to the network- driven bursts arising in the STG (Bucher et al. 2003). These spikes travel both antidromically and orthodromically, altering the output from the PD neurons and contributing to a tonic contraction of the PD muscles. Using intra-axonal voltage clamp recording, Ballo et  al. (2010) showed that DA enhances axonal I h through a direct cAMP-dependent mechanism, causing a prolonged depolarization that underlies the spike activity. These workers also demonstrated the remarkable dynamic properties of the axons, which alter PD spike amplitude, duration, and propagation velocity due to the interactions of multiple axonal currents (Ballo and Bucher 2009). Another site where ectopic spike initiation occurs is along the stn, the sole input nerve to the STG. A  number of regions along the stn show strong synaptotagmin-labeled neuropilar structures, and have terminals with dense-core vesicles, reminiscent of neurohemal release sites (Skiebe and Wollenschläger 2002). At one of these regions, where the stn branches with the two superior oesophageal nerves (sons), local application of octopamine evokes ectopic axonal spikes from the projection neuron MCN5 (Goaillard et al. 2004), which could affect the pyloric rhythm in the STG, independently of its spike pattern evoked from the CoG. As another example, the AGR neuron normally provides sensory input from the medial tooth protractor muscle to regulate the gastric mill and pyloric motor patterns (Combes et al. 1995). In the crab Cancer pagurus, it has an additional, anterior SIZ that generates tonic spiking in the dissected STNS, independent



Modulation of Crustacean Networks for Behavior

of sensory drive. Octopamine excites this anterior SIZ in a state-dependent manner, maximally increasing AGR tonic spiking in weakly active preparations (Daur et al. 2009). This is sufficient to enhance both the gastric mill and pyloric rhythms. The detailed pattern of spiking during a burst can also be modulated, which is important when the exact pattern of spike timing is the code rather than the smoothed average spike frequency. Selverston and colleagues have shown that the pattern of LP and PD bursting interspike intervals (ISIs) becomes more regular during DA application (Szucs et al. 2005), so that the timing of each spike over a sequence of bursts becomes more reproducible. The spike timing of the modulatory/sensory GPR neuron during an applied muscle stretch is affected by its own transmitters: allatostatin decreases both spike response rate and spike timing jitter, in part by increasing the membrane conductance, while 5HT has exactly the opposite effect (Billimoria et al. 2006). The sensory AGR neuron normally responds to muscle stretch by an increase in tonic spiking; however, neuropeptide F1 (TNRNFLRFamide) converts this into a bursting output, even in the absence of muscle stretch. Stretching the muscle changes the burst duration and spike frequency, which imparts a very different spike code from the resting state (Combes et al. 1997). These studies show that the detailed spike timing code is subject to modulation.

Modulation of Synaptic Interactions In addition to modulation of neuronal intrinsic firing properties, a second major mechanism for network reconfiguration is to modulate the strengths of the synapses in the network. The consequence of this modulation is to quantitatively “rewire” the network so the same anatomically defined network can generate a variety of functionally different circuits, each driving a modification on the basic behavior. The most detailed studies of synaptic modulation in the STNS have been performed with DA, 5HT, and octopamine in P. interruptus. Johnson and colleagues used a combination of photoinactivation and pharmacological blockade to isolate pairs of synaptically coupled neurons in the pyloric network, and determined the direct effects of each amine on each synapse in the entire network (Johnson and Harris-Warrick 1990, Johnson et al. 1993a, 1993b, 1994, 1995, Harris-Warrick and Johnson 2010) (Fig. 5.5). Several major conclusions were made. First, almost every synapse in the network was modified by each amine: there was no single major or critical target of amine action. This resembles the almost universal amine modulation of neuronal properties, described above. Second, a single amine has different and even opposing effects on different synapses in the network (Fig. 5.5A). For example, DA can strengthen some synapses while weakening others. Several functional synapses are silenced by DA. The output synapses of the cholinergic PD neurons become nonfunctional: the postsynaptic neurons remain sensitive to cholinergic agonists, suggesting that the PD terminals stop releasing transmitter. In other cases, a synapse may be functionally silent until the appropriate modulator is present. For example, the PY→LP chemical synapse is undetectable in the absence of modulatory inputs, but is activated by DA. Third, amines can strengthen or weaken electrical synapses as well as chemical synapses. In some cases, the sign of the change varies with the direction of current flow. For example, at the nonrectifying AB-PD electrical synapse, DA enhances coupling strength in the PD→AB direction, but weakens it in the AB→PD direction. This curious result arises from differential effects of DA on the input resistances of the two neurons, affecting current flow from the electrodes or synaptic input sites to the gap junctions (Johnson et al. 1993a). Fourth, some pyloric neurons are coupled by both electrical and chemical synapses; these can be independently modulated, resulting in a variety of different synaptic interactions, and even inverting the

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Fig. 5.5. Modulation of synaptic interactions in the stomatogastric ganglion. (A) Summary of effects of DA on selected synapses in the pyloric network. Graded transmission is measured in the presence of tetrodotoxin. In each panel, the bottom neuron is depolarized and the synaptic response of the top neuron shown. Synapses are strengthened, weakened, activated, and abolished, and even inverted in sign when DA is added (from Harris-Warrick et al. 1998, with permission from John Wiley and Sons). (B) Summary of the presynaptic and postsynaptic effects of DA in the pyloric network. Bolded lines indicate synapses that are strengthened by DA, and dashed lines indicate synapses that are weakened by DA. Effects of DA on input resistance are indicated by circled symbols within each neuron. Shaded circles show synapses where the pre- and postsynaptic effects of DA are of opposite functional sign. Effects of DA on neuronal responses to glutamate iontophoresis are shown in the pipette symbols next to each neuron. Effects of DA on transmitter release and Ca 2+ accumulation are indicated in the terminal symbol for synapses. Question mark indicates uncertain functional connection (from Harris-Warrick and Johnson 2010, with permission from the authors). (C) RPCH strengthens the LP→PD synapse during the ongoing pyloric cycle. (D) RPCH dramatically enhances graded synapse from LP to PD neurons. (E) Despite strengthening of the LP-PD synapses, which is the only feedback inhibitory synapse to the pyloric pacemaker kernel, RPCH does not alter the ability of LP to reset the cycle. Phase response curve shows the same Pacemaker response to LP stimulation under control conditions (open circles) and during RPCH (filled circles). C–E from Thirumalai et al. 2006, with permission from The American Physiological Society. Neuron names: AB, anterior burster; LP, lateral pyloric, PD, pyloric dilator; PY, pyloric constrictor; VD, ventricular dilator.



Modulation of Crustacean Networks for Behavior

net sign of the synapse (Johnson et al. 1993b, 1994). For example, in the absence of modulators, the LP→PY synapse is depolarizing, reflecting a functional electrical synapse while the chemical synapse is silent. Dopamine converts this synapse to a hyperpolarizing one, as the electrical synapse becomes weaker and the silent inhibitory chemical synapse is strongly activated (Fig. 5.5A). Serotonin and octopamine can also invert mixed synapses in the pyloric network (Johnson et al. 1994). Fifth, the synapses in the pyloric network function by a combination of spike-evoked release and graded voltage-dependent release (Hartline and Graubard 1992), and these two synaptic components can be independently modulated (Ayali et al. 1998). For example, at three synapses in the pyloric network, DA enhances graded transmission, but has different effects on spike-evoked transmission depending on DA’s effects on the postsynaptic neuron’s input resistance. At two of these, the graded synaptic component was enhanced but the spike-evoked component was weakened. Thus, the functional effect of the modulator will depend on the relative importance of the graded versus spike-evoked component of the synapses. Although less work has been done with other modulators, they also affect synaptic transmission. For example, the peptide RPCH has complex effects on the pyloric rhythm due to its interactions with the cardiac sac network (Dickinson et al. 2001). Also, RPCH greatly strengthens the synaptic connections from the modulatory IVN neurons onto pyloric neurons, which alters cardiac sac-pyloric interactions and causes the pyloric VD neuron to switch to the cardiac sac pattern. Mechanisms of Synaptic Modulation A neuromodulator can alter synaptic strength by presynaptic actions to alter transmitter release, and/or by postsynaptic actions to alter responsiveness to the transmitter. Kloppenburg et al. (2000, 2007) used calcium imaging to look for changes in presynaptic voltage-activated calcium accumulation in single pyloric neuron nerve terminals (Fig. 5.5B). In general, synapses strengthened by DA also showed increased voltage-dependent calcium accumulation in the presynaptic terminals, while weakened synapses showed reduced calcium accumulation. However, detailed study showed considerable variability in responses of individual terminals (Kloppenburg et al. 2007). In PD neurons, half of the terminals showed decreased voltage-dependent calcium accumulation with DA, consistent with a reduction in synaptic strength; however, the remaining terminals showed no change, even if they were on the same neuron. Comparable variability was seen in LP synapses. This variability may reflect the spatial nonhomogeneity of DA receptors on pyloric neuron dendrites, as recently described by Oginsky et al. (2010). The situation is even more complex in PY synapses (Kloppenburg et al. 2007). Electrophysiologically recorded PY output synapses are uniformly strengthened by DA. However, only 38% of PY terminals increased calcium accumulation with DA, while 20% significantly decreased calcium accumulation, in apparent opposition to the overall effect of DA. Paired optical recordings showed that these opposing responses could occur simultaneously in a single neuron. These results show the heterogeneity in modulatory responses in a neuron, which raises questions about the integrated responses to modulators in a network (see below). Postsynaptic mechanisms for synaptic modulation could reflect simple changes in input resistance, alterations in receptor number or sensitivity, or changes in voltage-dependent amplifying currents activated by the synaptic event. Johnson and Harris-Warrick (1997) monitored amine effects on several postsynaptic responses to iontophoresis of the synapse’s transmitter, glutamate (Fig. 5.5B). Many of these postsynaptic responses paralleled the amine’s effect on synaptic strength: when DA strengthened a synapse, it usually enhanced the postsynaptic response to transmitter. However, there were a number of synapses where the postsynaptic response was of the opposite sign to the recorded synaptic strength change. For example, DA enhanced the LP→PD synapse and increased presynaptic calcium accumulation in many LP nerve terminals; however,

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Ronald Harris-Warrick it reduced the postsynaptic PD response to LP’s transmitter, glutamate. Clearly, the presynaptic effect must outweigh the opposing postsynaptic effect to yield a net increase in synaptic strength. Functional Importance of Synaptic Modulation for Network Function The modulatory changes in synaptic strength described above could certainly play an important role in shaping the motor pattern generated by the modified network. However, in some cases, this has not been borne out. For example, the LP neuron provides the sole inhibitory feedback to the pyloric pacemaker kernel, and changes in the strength of this synapse should modify the cycle frequency. Thirumalai et al. (2006) found that RPCH potentiates this synapse but has no effect on cycle frequency, because the LP was firing during a phase when the pacemakers are already maximally hyperpolarized by their intrinsic currents and the additional synaptic inhibition had no further effect (Fig. 5.5C). Similar results have been seen with DA (Johnson et al. 2011), which strengthens this synapse, but constrains LP activity (by a combination of intrinsic effects and synaptic inhibition from PY neurons) to fire at a phase when the pacemakers are insensitive to input. In contrast, 5HT weakens the synapse, but through other effects shifts the phase of LP activity to enhance its influence on cycle frequency. This work shows that synaptic modulation must be analyzed in the full context of other effects of the modulator on intrinsic firing and network properties to determine its functional importance.

MODULATION OF THE NEUROMUSCULAR TRANSFORM Although the CPG networks generate the pattern of activity of motor neuron firing to drive behaviors, the transformation of the motor neuron pattern to muscle contraction is not a simple one. Muscles have many intrinsic properties that filter or alter the timing of the behavior as commanded by the motor neurons. The kinetics of their contractions can dramatically change the behavioral output. For example, in vivo, the pyloric pattern is periodically altered by interactions with the slower gastric mill and cardiac sac networks, which can subtly alter the spike frequency or duration in pyloric motor neuron bursts. When the effect of these subtle alterations in motor pattern reach the pyloric muscles, some of them have sufficiently slow contraction/relaxation kinetics that their major movement is timed with the cardiac sac and gastric mill rhythms, with only subtle effects of the pyloric rhythm (Thuma et al. 2003). This neuromuscular transform is subject to neuromodulation, which will change the behavior in additional ways beyond modulation of the CPG output. The details of modulation of muscle and the neuromuscular junctions in crustaceans are discussed in the c­ hapters 4 and 6 by Atwood and Lnenicka, respectively. Monoamines and peptides can directly evoke pyloric muscle contraction (Lingle 1981)  as well as modulating transmitter release from the motor neuron terminals and the muscle response to released transmitter. In the shrimp P.  serratus, FMRFamide evokes myogenic oscillations in PD muscles (Meyrand and Marder 1991); this results in a complex interaction of the myogenic and neurogenic rhythms, where the neural input serves primarily as a trigger to evoke all-or-none muscle contractions.

CONCLUSIONS AND FUTURE DIRECTIONS Due to the relative numerical simplicity of crustacean neural networks, we know more about their function and modulation than any vertebrate network and most other invertebrate networks. This short review has described a number of general principles, leading to many



Modulation of Crustacean Networks for Behavior

questions for future research. While they arise from studies of crustacean neuromodulation, they are generic and can be applied to networks of all kinds in all animals. Sources of Neuromodulators 1. The complexity of modulatory inputs to crustacean networks is enormous, and it is not clear why so many neuromodulators, many with multiple mechanisms of action, are necessary. Nearly every known transmitter substance can act as a neuromodulator; the sole current exception is glycine, which appears to act only via ionotropic receptors. The number of neuropeptide families, and the number of members in each family, continues to grow with new biochemical tools for their detection. Why are there so many different peptides? Do they all play different roles or do they converge onto a few basic mechanisms, as in the STG? A number of atypical transmitters have been identified, including gases such as NO and carbon monoxide (Mann and Motterlini 2007), fatty acid derivatives such as arachidonic acid and the endocannabinoids (Katona and Freund 2012), and N-acyl-conjugated neurotransmitters (Connor et al. 2010). These have not yet been tested in any crustacean system, so the complexity of modulatory action could be still greater. Understanding this complexity will depend on more careful identification of the receptors that neuromodulators activate. For example, is the apparent convergence of peptide actions in the STG due to a small number of homologous peptide receptors that all activate the same second messenger systems, while the divergence of monoamine actions results from activation of multiple receptor types for each amine? 2. Neuromodulators act over a spectrum of spatial and temporal specificity, ranging from local transmitter release to volume transmission to hormonal actions that affect the entire body. A  single peptide or monoamine may affect a network via several pathways. How are these different actions coordinated to achieve maximal effect in regulating network function in the intact animal? Are there separate receptors to mediate the hormonal and the transmitter actions of a modulator, which act over very different concentration ranges? Many studies of modulator action have used bath application, which mimics hormonal actions, and to a lesser extent, volume transmission, but lacks the spatial and temporal specificity when the compound is released by neuronal firing activity. It will be important to determine whether the earlier studies accurately reflect the endogenous actions of the modulator in vivo. 3. Modulatory projection neurons typically release multiple transmitters, often combining a small transmitter with one or more peptides. Thus, to mimic the actions of these peptides, the interactions between the cotransmitters will have to be carefully studied, rather than each transmitter in isolation. Some neurons do not respond to all of the cotransmitters of a projection neuron: does this result from different spatial localization of the various receptors relative to the release sites or actions of extracellular peptidases and other enzymes? These questions are just starting to be addressed. 4. There are many modulatory neurons for each network. In the crab STNS, around 40 neurons modify STG networks. How is the activity of these neurons coordinated? Is there a metamodulatory circuitry that regulates the activity of these projection neurons (Stein 2009)? How is this circuitry organized and controlled to determine the appropriate pattern of modulator activity for behavior? Most studies of modulatory projection neurons have studied just one at a time, but in vivo it is likely that many of them are coactive at any moment. What are the consequences of this coactivation? Network Modulation 5. Behavioral networks have evolved for maximal flexibility, to allow a single anatomically defined network to generate many variants on a basic motor theme. In part, this is regulated by modulatory

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Ronald Harris-Warrick inputs. Neuromodulators can select which component neurons are active in the network; they can modify the intrinsic firing properties and responsiveness of the network neurons in many different ways, and they can alter the strengths of network synapses to functionally “rewire” the network for a particular behavior. They can also combine separate networks into larger conjoint networks to organize more complex behaviors. Some neural networks, such as those in the STG, require modulatory inputs to function at all. Are these modulatory inputs the switches that normally activate the behavior, or do they prepare the network for activation by more rapid transmitter switches? 6. Modulation can be both extrinsic and intrinsic to a neural network, and the consequences for network function are very different in the two cases. Intrinsic neuromodulation occurs whenever the network is functioning and forms another potential target for extrinsic modulation. How are these two forms of modulation coordinated in the intact network? 7. Central and peripheral modulatory actions must be coordinated so that the muscle and sensory responses are appropriate to the central motor pattern generated by the neural network. Significant progress has been made on this problem in the crustacean heart (Fort et al. 2004) and in the Aplysia feeding network (Brezina et al. 2000), but the detailed mechanisms of this coordination are still unknown. This will be essential before we begin to understand the actions of neuromodulators in vivo, which will combine the separate actions at many sites to generate appropriate behaviors. Cellular and Synaptic Mechanisms of Network Modulation 8. Neuromodulators change the baseline intrinsic electrophysiological properties of network neurons in many ways. They can activate or inhibit rhythmic bursting, bistability, postinhibitory rebound, and delayed excitation and can regulate spike frequency to shape the activity of each neuron in the motor pattern. These effects are mediated in part by modulation of the biophysical properties of ion currents in the neurons, including their maximal conductance, voltage dependence, and kinetics. Different neuromodulators can evoke very similar activity patterns by very different ionic mechanisms, supporting the general principle that multiple ionic mechanisms can generate an activity pattern (Prinz et al. 2004). Neuromodulators can also regulate the regularity of spike timing during a burst, which may have important consequences for the pattern of movement as filtered through the muscle contraction kinetics. 9. While some modulators have a fairly simple action to excite a neuron by activating one current, others affect a variety of different currents in a single cell. The significance of this result is unclear. At the level of the neuron, which ionic current changes are most important for shaping the neuron’s activity, and which play subordinate roles or perhaps are not relevant to the neuron’s activity except in specific contexts? 10. Modulators can shape the synaptic interactions between network neurons by both presynaptic regulation of transmitter release and postsynaptic regulation of responsiveness to the transmitter. These can be mediated by different mechanisms at different synapses, and may reflect global changes in input resistance, or terminal-specific changes in calcium currents. In crustaceans and other invertebrates, many neurons use both spike-evoked and graded transmission at synapses, and these can be differentially modulated to shape the synaptic response. What is the significance of all this synaptic flexibility for the function of the network? 11. Neuromodulators can evoke opposing effects on a neuron or a synapse, with some actions apparently opposing the net effect of the modulation (Harris-Warrick and Johnson 2010). Why would this occur? Does it arise simply from evolutionary noise, where “mistakes” are tolerated so long as they do not block the overall modulatory effect? Or does this reflect a state-dependent effect of the neuromodulator, which can reverse its effect under the right conditions? Or do these



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opposing actions constrain the flexibility of the network, preventing it from being overmodulated and becoming nonfunctional? 12. Modulator actions can only be understood from the sum of their actions throughout the network and the animal, and not from single actions identified at a specific synapse or neuron. In several cases in the STG, the functional consequences of synaptic modulation have been negated by other modulatory actions elsewhere in the network. While these are extreme examples, the effects of modulatory events at any one point in a system are always regulated by events at all other points in the system. This major point has not been studied in any detail in any system and poses a major challenge for future research into the neural bases of behavior. In conclusion, while the complexity of neural network modulation seems almost overwhelming, it makes sense to the animal and to its nervous system. These systems have evolved for a purpose, to ensure maximal behavioral flexibility for all situations, and are essential for normal behavior. Due to their relative numerical simplicity and accessibility, the crustacean networks will continue to be prime models for studying the role and mechanisms of behavioral modulation for many years to come.

ACKNOWLEDGMENTS This work was supported by NIH grant NS17323.

REFERENCES Ayali, A., and R.M. Harris-Warrick. 1999. Monoamine control of the pacemaker kernel and cycle frequency in the lobster pyloric network. Journal of Neuroscience 19:6712–6722. Ayali, A., B.R. Johnson, and R.M. Harris-Warrick. 1998. Dopamine modulates graded and spike evoked synaptic inhibition independently at single synapses in the pyloric network of the lobster. Journal of Neurophysiology 79:2063–2069. Ballo, A.W., and D. Bucher. 2009. Complex intrinsic membrane properties and dopamine shape spiking activity in a motor axon. Journal of Neuroscience 29:5062–5074. Ballo, A.W., J.C. Keene, P.J. Troy, M.L. Goeritz, F. Nadim, and D. Bucher. 2010. Dopamine modulates Ih in a motor axon. Journal of Neuroscience 30:8425–8434. Beltz, B., J.S. Eisen, R. Flamm, R. Harris-Warrick, S. Hooper, and E. Marder. 1984. Serotonergic innervation and modulation of the stomatogastric ganglion of three decapod crustaceans (Panulirus interruptus, Homarus americanus, and Cancer irroratus). Journal of Experimental Biology 109:35–54. Benson, J.A., and I.M. Cooke. 1984. Driver potentials and the organization of rhythmic bursting in crustacean ganglia. Trends in Neuroscience 7:85–91. Billimoria, C.P., R.A. Dicaprio, J.T. Birmingham, L.F. Abbott, and E. Marder. 2006. Neuromodulation of spike-timing precision in sensory neurons. Journal of Neuroscience 26:5910–5919. Birmingham, J.T., C.P. Billimoria, T.R. Deklotz, R.A. Stewart, and E. Marder. 2003. Differential and history-dependent modulation of a stretch receptor in the stomatogastric system of the crab, Cancer borealis. Journal of Neurophysiology 90:3608–3616. Blitz, D.M., M.P. Beenhakker, and M.P. Nusbaum. 2004. Different sensory systems share projection neurons but elicit distinct motor patterns. Journal of Neuroscience 24:11381–11390. Braun, G., and B. Mulloney. 1993. Cholinergic modulation of the swimmeret motor system in crayfish. Journal of Neurophysiology 70:2391–2398. Brezina, V., I.V. Orekhova, and K.R. Weiss. 2000. Optimization of rhythmic behaviors by modulation of the neuromuscular transform. Journal of Neurophysiology 83:260–279. Bucher, D., V. Thirumalai, and E. Marder. 2003. Axonal dopamine receptors activate peripheral spike initiation in a stomatogastric motor neuron. Journal of Neuroscience 23:6866–6875. Cebada, J., and U. García. 2007. Histamine operates Cl--gated channels in crayfish neurosecretory cells. Journal of Experimental Biology 210:3962–3969.

139

140

Ronald Harris-Warrick Chen, R., S.S. Cape, R.M. Sturm, and L. Li. 2010a. Mass spectrometric imaging of neuropeptides in decapod crustacean neuronal tissues. Methods in Molecular Biology 656:451–463. Chen, R., L. Hui, S.S. Cape, J. Wang, and L. Li. 2010b. Comparative neuropeptidomic analysis of food intake via a multi-faceted mass spectrometric approach. ACS Chemical Neuroscience 1:204–214. Christie, A.E., W. Stein, J.E. Quinlan, M.P. Beenhakker, E. Marder, and M.P. Nusbaum. 2004. Actions of a histaminergic/peptidergic projection neuron on rhythmic motor patterns in the stomatogastric nervous system of the crab Cancer borealis. Journal of Comparative Neurology 469:153–169. Christie, A.E., E.A. Stemmler, and P.S. Dickinson. 2010a. Crustacean neuropeptides. Cellular and Molecular Life Sciences 67:4135–4169. Christie, A.E., J.S. Stevens, M.R. Bowers, M.C. Chapline, D.A. Jensen, K.M. Schegg, J. Goldwaser, M.A. Kwiatkowski, T.K. Pleasant, Jr., L. Shoenfeld, L.K. Tempest, C.R. Williams, T. Wiwatpanit, C.M. Smith, K.M. Beale, D.W. Towle, D.A. Schooley, and P.S. Dickinson. 2010b. Identification of a calcitonin-like diuretic hormone that functions as an intrinsic modulator of the American lobster, Homarus americanus, cardiac neuromuscular system. Journal of Experimental Biology 213:118–127. Claiborne, B.J., and A.I. Selverston. 1984. Histamine as a neurotransmitter in the stomatogastric nervous system of the spiny lobster. Journal of Neuroscience 4:708–721. Clark, M.C., and D.J. Baro. 2006. Molecular cloning and characterization of crustacean type-one dopamine receptors: D1α Pan and D1βPan. Comparative Biochemistry and Physiology 143B:294–301. Clark, M.C., and D.J. Baro. 2007. Arthropod D2 receptors positively couple with cAMP through the Gi/o protein family. Comparative Biochemistry and Physiology 146B: 9–19. Clark, M.C., T.E. Dever, J.J. Dever, P. Xu, V. Rehder, M.A. Sosa, and D.J. Baro. 2004. Arthropod 5-HT2 receptors: a neurohormonal receptor in decapod crustaceans that displays agonist independent activity resulting from an evolutionary alteration to the DRY motif. Journal of Neuroscience 24:3421–3435. Clark, M.C., R. Khan, and D.J. Baro. 2008. Crustacean dopamine receptors: localization and G protein coupling in the stomatogastric ganglion. Journal of Neurochemistry 104:1006–1019. Coleman, M.J., M.P. Nusbaum, I. Cournil, and B.J. Claiborne. 1992. Distribution of modulatory inputs to the stomatogastric ganglion of the crab, Cancer borealis. Journal of Comparative Neurology 325:581–594. Combes, D., J. Simmers, and M. Moulins. 1995. Structural and functional characterization of a muscle tendon proprioceptor in lobster. Journal of Comparative Neurology 363:221–234. Combes, D., J. Simmers, and M. Moulins. 1997. Conditional dendritic oscillators in a lobster mechanoreceptor neurone. Journal of Physiology (London) 499:161–177. Connor, M., C.W. Vaughan, and R.J. Vandenberg. 2010. N-acyl amino acids and N-acyl neurotransmitter conjugates: neuromodulators and probes for new drug targets. British Journal of Pharmacology 160:1857–1871. Cooke, I.M. 1966. The sites of action of pericardial organ extract and 5-hydroxytryptamine in the decapod crustacean heart. American Zoologist 6:107–121. Cooke, I.M. 2002. Reliable, responsive pacemaking and pattern generation with minimal cell numbers: the crustacean cardiac ganglion. Biological Bulletin 202:108–136. Cropper, E. C., P.E. Lloyd, W. Reed, R. Tenenbaum, I. Kupfermann, and K.R. Weiss. 1987. Multiple neuropeptides in cholinergic motor neurons of Aplysia: evidence for modulation intrinsic to the motor circuit. Proceedings of the National Academy of Sciences of the United States of America 84:3486–3490. Cruz-Bermúdez, N.D., and E. Marder. 2007. Multiple modulators act on the cardiac ganglion of the crab, Cancer borealis. Journal of Experimental Biology 210:2873–2884. Daur, N., F. Nadim, and W. Stein. 2009. Regulation of motor patterns by the central spike-initiation zone of a sensory neuron. European Journal of Neuroscience 30:808–822. Delong, N.D., M. P. Beenhakker, and M.P. Nusbaum. 2009. Presynaptic inhibition selectively weakens peptidergic cotransmission in a small motor system. Journal of Neurophysiology 102:3492–3504. Delong, N.D., and M.P. Nusbaum. 2010. Hormonal modulation of sensorimotor integration. Journal of Neuroscience 30:2418–2427. Dickinson, P.S. 1995. Interactions among neural networks for behavior. Current Opinion in Neurobiology 5:792–798.



Modulation of Crustacean Networks for Behavior

Dickinson, P.S., and M. Moulins. 1992. Interactions and combinations between different networks in the stomatogastric nervous system. Pages 139–160 in R.M. Harris-Warrick, E. Marder, A.I. Selverston, and M. Moulins, editors. Dynamic biological networks: the stomatogastric nervous system. MIT Press, Cambridge. Dickinson, P.S., C. Mecsas, and E. Marder. 1990. Neuropeptide fusion of two motor-pattern generator circuits. Nature 344:155–158. Dickinson, P.S., W.P. Fairfield, J.R. Hetling, and J. Hauptman. 1997. Neurotransmitter interactions in the stomatogastric system of the spiny lobster: one peptide alters the response of a central pattern generator to a second peptide. Journal of Neurophysiology 77:599–610. Dickinson, P.S., J. Hauptman, J. Hetling, and A. Mahadevan. 2001. RPCH modulation of a multi-oscillator network: effects on the pyloric network of the spiny lobster. Journal of Neurophysiology 85:1424–1435. Dickinson, P.S., E.A. Stemmler, and A.E. Christie. 2008. The pyloric neural circuit of the herbivorous crab Pugettia producta shows limited sensitivity to several neuromodulators that elicit robust effects in more opportunistically feeding decapods. Journal of Experimental Biology 211:1434–1447. El Manira, A., and F. Clarac. 1994. Presynaptic inhibition is mediated by histamine and GABA in the crustacean escape reaction. Journal of Neurophysiology 71:1088–1095. Elson, R.C., and A.I. Selverston. 1997. Evidence for a persistent Na+ conductance in neurons of the gastric mill rhythm generator of spiny lobsters. Journal of Experimental Biology 200:1795–1807. Faumont, S., D. Combes, P. Meyrand, and J. Simmers. 2005. Reconfiguration of multiple motor networks by short- and long-term actions of an identified modulatory neuron. European Journal of Neuroscience 22:2489–2502. Flamm, R.E., and R.M. Harris-Warrick. 1986. Aminergic modulation in the lobster stomatogastric ganglion. II. Target neurons of dopamine, octopamine and serotonin within the pyloric circuit. Journal of Neurophysiology 55:866–881. Florey, E. 1967. Neurotransmitters and modulators in the animal kingdom. Federation Proceedings 26:1164–1178. Florey, E., and M. Rathmayer. 1978. The effects of octopamine and other amines on the heart and on neuromuscular transmission in decapod crustaceans: further evidence for a role as neurohormone. Comparative Biochemistry and Physiology 61C:229–237. Fort, T.J., V. Brezina, and M.W. Miller. 2004. Modulation of an integrated central pattern generator-effector system: dopaminergic regulation of cardiac activity in the blue crab Callinectes sapidus. Journal of Neurophysiology 92:3455–3470. Freschi, J.E. 1989. Proctolin activates a slow, voltage-dependent sodium current in motoneurons of the lobster cardiac ganglion. Neuroscience Letters 106:105–111. Freschi, J.E., and D.R. Livengood. 1989. Membrane current underlying muscarinic cholinergic excitation of motoneurons in lobster cardiac ganglion. Journal of Neurophysiology 62:984–995. Fu, Q., K.K. Kutz, J.J. Schmidt, Y.W. Hsu, D.I. Messinger, S.D. Cain, H.O. De La Iglesia, A.E. Christie, and L. Li. 2005. Hormone complement of the Cancer productus sinus gland and pericardial organ: an anatomical and mass spectrometric investigation. Journal of Comparative Neurology 493:607–626. Goaillard, J.M., D.J. Schulz, V.L. Kilman, and E. Marder. 2004. Octopamine modulates the axons of modulatory projection neurons. Journal of Neuroscience 24:7063–7073. Gola, M., and A. Selverston. 1981. Ionic requirements for bursting activity in lobster stomatogastric neurons. Journal of Comparative Physiology A 145:191–207. Golowasch, J., and E. Marder. 1992. Proctolin activates an inward current whose voltage-dependence is modified by extracellular Ca 2+. Journal of Neuroscience 12:810–817. Goy, M.F. 2005. Nitric oxide: an inhibitory retrograde modulator in the crustacean heart. Comparative Biochemistry and Physiology 142A:151–163. Grundfest, H., and J.P. Reuben. 1961. Neuromuscular synaptic activity in lobster. Pages 92–104 in E. Florey, editor. Nervous inhibition. Pergamon Press, Oxford. Harris-Warrick, R.M. 1988. Chemical modulation of central pattern generators. Pages 285–331 in A.H. Cohen, S. Rossignol, and S. Grillner, editors. Neural control of rhythmic movements. John Wiley & Sons, New York.

141

142

Ronald Harris-Warrick Harris-Warrick, R.M., and R.E. Flamm. 1986. Chemical modulation of a small central pattern generator circuit. Trends in Neuroscience 9:432–437. Harris-Warrick, R.M., and R.E. Flamm. 1987. Multiple mechanisms of bursting in a conditional bursting neuron. Journal of Neuroscience 7:2113–2128. Harris-Warrick, R.M., and B.R. Johnson. 2010. Checks and balances in neuromodulation. Frontiers in Behavioral Neuroscience 4:47. Harris-Warrick, R.M., and E.A. Kravitz. 1984. Cellular mechanisms for modulation of posture by serotonin and octopamine in the lobster. Journal of Neuroscience 4:1976–1993. Harris-Warrick, R.M., F. Nagy, and M.P. Nusbaum. 1992. Neuromodulation of stomatogastric networks by identified neurons and transmitters. Pages 87–138 in R.M. Harris-Warrick, E. Marder, A.I. Selverston, and M. Moulins, editors. Dynamic biological networks: the stomatogastric nervous system. MIT Press, Cambridge, MA. Harris-Warrick, R.M., L.M. Coniglio, N. Barazangi, J. Guckenheimer, and S. Gueron. 1995a. Dopamine modulation of transient potassium current evokes phase shifts in a central pattern generator network. Journal of Neuroscience 15:342–358. Harris-Warrick, R.M., L.M. Coniglio, R.M. Levini, S. Gueron, and J. Guckenheimer. 1995b. Dopamine modulation of two subthreshold currents produces phase shifts in activity of an identified motoneuron. Journal of Neurophysiology 74:1404–1420. Harris-Warrick, R.M., B.R. Johnson, J.H. Peck, P. Kloppenburg, A. Ayali, and J. Skarbinski. 1998. Distributed effects of dopamine modulation in the crustacean pyloric network. Annals of the New York Academy of Sciences 860:155–167. Hartline, D.K., and K. Graubard. 1992. Cellular and synaptic properties in the crustacean stomatogastric nervous system. Pages 31–86 in R.M. Harris-Warrick, E. Marder, A.I. Selverston, and M. Moulins, editors. Dynamic biological networks: the stomatogastric nervous system. MIT Press, Cambridge, MA. Hashemzadeh-Gargari, H., and J.E. Freschi. 1992. Histamine activates chloride conductance in motor neurons of the lobster cardiac ganglion. Journal of Neurophysiology 68:9–15. Hedrich, U.B., and W. Stein. 2008. Characterization of a descending pathway: activation and effects on motor patterns in the brachyuran crustacean stomatogastric nervous system. Journal of Experimental Biology 211:2624–2637. Hooper, S.L., and E. Marder. 1987. Modulation of the lobster pyloric rhythm by the peptide proctolin. Journal of Neuroscience 7:2097–2112. Hooper, S.L., and M. Moulins. 1990. Cellular and synaptic mechanisms responsible for a long-lasting restructuring of the lobster pyloric network. Journal of Neurophysiology 64:1574–1589. Hsieh, S.L., S.M. Chen, Y.H. Yang, and C.M. Kuo. 2006. Involvement of norepinephrine in the hyperglycemic responses of the freshwater giant prawn, Macrobrachium rosenbergii, under cold shock. Comparative Biochemistry and Physiology 143A:254–263. Johnson, B.R., and R.M. Harris-Warrick. 1990. Aminergic modulation of graded synaptic transmission in the lobster stomatogastric ganglion. Journal of Neuroscience 10:2066–2076. Johnson, B.R., and R.M. Harris-Warrick. 1997. Amine modulation of glutamate responses from pyloric motor neurons in the lobster stomatogastric ganglion. Journal of Neurophysiology 78:3210–3221. Johnson, B.R., J.H. Peck, and R.M. Harris-Warrick. 1993a. Amine modulation of electrical coupling in the pyloric network of the lobster stomatogastric ganglion. Journal of Comparative Physiology A 172:715–732. Johnson, B.R., J.H. Peck, and R.M. Harris-Warrick. 1993b. Dopamine induces sign reversal at mixed chemical-electrical synapses. Brain Research 625:159–164. Johnson, B.R., J.H. Peck, and R.M. Harris-Warrick. 1994. Differential modulation of chemical and electrical components of mixed synapses in the lobster stomatogastric ganglion. Journal of Comparative Physiology A 175:233–249. Johnson, B.R., J.H. Peck, and R.M. Harris-Warrick. 1995. Distributed amine modulation of graded chemical transmission in the pyloric network of the lobster stomatogastric ganglion. Journal of Neurophysiology 74:437–452.



Modulation of Crustacean Networks for Behavior

Johnson, B.R., J.M. Brown, M.D. Kvarta, J.Y. Lu, L.R. Schneider, F. Nadim, and R.M. Harris-Warrick. 2011. Differential modulation of synaptic strength and timing regulate synaptic efficacy in a motor network. Journal of Neurophysiology 105:293–304. Kadiri, L.R., A.C. Kwan, W.W. Webb, and R.M. Harris-Warrick. 2011. Dopamine-induced oscillations of the pyloric pacemaker neuron rely on release of calcium from intracellular stores. Journal of Neurophysiology 106:1288–1298. Katona, I., and T.F. Freund. 2012. Multiple functions of endocannabinoid signaling in the brain. Annual Review of Neuroscience 35:529–558. Katz, P.S. 1989. Motor pattern modulation by serotonergic sensory cells in the stomatogastric nervous system. PhD thesis, Cornell University, Ithaca, NY. Katz, P.S. 1995. Intrinsic and extrinsic neuromodulation of motor circuits. Current Opinion in Neurobiology 5:799–808. Katz, P.S. 1999. Beyond neurotransmission: neuromodulation and its importance for information processing. Oxford University Press, Oxford, United Kingdom. Katz, P.S., and R.M. Harris-Warrick. 1989. Serotonergic/cholinergic muscle receptor cells in the crab stomatogastric nervous system. II. Rapid nicotinic and prolonged modulatory effects on neurons in the stomatogastric ganglion. Journal of Neurophysiology 62:571–581. Katz, P.S., and R.M. Harris-Warrick. 1990. Neuromodulation of the crab pyloric central pattern generator by serotonergic/cholinergic proprioceptive afferents. Journal of Neuroscience 10:1495–1512. Katz, P.S., M.H. Eigg, and R.M. Harris-Warrick. 1989. Serotonergic/cholinergic muscle receptor cells in the crab stomatogastric nervous system. I. Identification and characterization of the gastropyloric receptor cells. Journal of Neurophysiology 62:558–570. Kiehn, O., and R.M. Harris-Warrick. 1992a. 5-HT modulation of hyperpolarization-activated inward current and calcium-dependent outward current in a crustacean motor neuron. Journal of Neurophysiology 68:496–508. Kiehn, O., and R.M. Harris-Warrick. 1992b. Serotonergic stretch receptors induce plateau properties in a crustacean motor neuron by a dual-conductance mechanism. Journal of Neurophysiology 68:485–495. King, D.G. 1976. Organization of crustacean neuropil. I. Patterns of synaptic connections in lobster stomatogastric ganglion. Journal of Neurocytology 5:207–237. Kloppenburg, P., W.R Zipfel, W.W. Webb, and R.M. Harris-Warrick. 2000. Highly localized Ca 2+ accumulation revealed by multiphoton microscopy in an identified motoneuron, and its modulation by dopamine. Journal of Neuroscience 20:2523–2533. Kloppenburg, P., W.R. Zipfel, W.W. Webb, and R.M. Harris-Warrick. 2007. Heterogeneous effects of dopamine on highly localized, voltage-induced Ca 2+ accumulation in identified motoneurons. Journal of Neurophysiology 98:2910–2917. Kravitz, E.A., S. Glusman, R.M. Harris-Warrick, M.Livingstone, T. Schwarz, and M.F. Goy. 1980. Amines and a peptide as neurohormones in lobsters: actions on neuromuscular preparations and preliminary behavioral studies. Journal of Experimental Biology 89:159–176. Krenz, W.D., D. Nguyen, N.L. Pérez-Acevedo, and A.I. Selverston. 2000. Group I, II, and III mGluR compounds affect rhythm generation in the gastric circuit of the crustacean stomatogastric ganglion. Journal of Neurophysiology 83:1188–1201. Kupfermann, I. 1979. Modulatory actions of neurotransmitters. Annual Review of Neuroscience 2:447–465. Lingle, C. 1981. The modulatory action of dopamine on crustacean foregut neuromuscular preparations. Journal of Experimental Biology 94:285–299. Livingstone, M.S., R.M. Harris-Warrick, and E.A. Kravitz. 1980. Serotonin and octopamine produce opposite postures in lobsters. Science 208:76–79. Mann, B.E., and R. Motterlini. 2007. CO and NO in medicine. Chemical Communications:4197–4208. Marder, E. 1974. Acetylcholine as an excitatory neuromuscular transmitter in the stomatogastric system of the lobster. Nature 251:730–731. Marder, E., and D. Bucher. 2007. Understanding circuit dynamics using the stomatogastric nervous system of lobsters and crabs. Annual Review of Physiology 69:291–316.

143

144

Ronald Harris-Warrick Marder, E., and V. Thirumalai. 2002. Cellular, synaptic and network effects of neuromodulation. Neural Networks 15:479–493. Meyrand, P., and E. Marder. 1991. Matching neural and muscle oscillators: control by FMRFamide-like peptides. Journal of Neuroscience 11:1150–1161. Meyrand, P., J. Simmers, and M. Moulins. 1991. Construction of a pattern-generating circuit with neurons of different networks. Nature 351:60–63. Meyrand, P., J. Simmers, and M. Moulins. 1994. Dynamic construction of a neural network from multiple pattern generators in the lobster stomatogastric nervous system. Journal of Neuroscience 14:630–644. Mulloney, B., and C. Smarandache-Wellmann. 2012. Neurobiology of the crustacean swimmeret system. Progress in Neurobiology 96:242–267. Mulloney, B., L.D. Acevedo, and A.G. Bradbury. 1987. Modulation of the crayfish swimmeret rhythm by octopamine and the neuropeptide proctolin. Journal of Neurophysiology 58:584–597. Mulloney, B., H. Namba, H.J. Agricola, and W.M. Hall. 1997. Modulation of force during locomotion: differential action of crustacean cardioactive peptide on power-stroke and returnstroke motor neurons. Journal of Neuroscience 17:6872–6883. Nagy, F., and P. S. Dickinson. 1983. Control of a central pattern generator by an identified modulatory interneurone in crustacea. I. Modulation of the pyloric motor output. Journal of Experimental Biology 105:33–58. Nusbaum, M.P., and M.P. Beenhakker. 2002. A small-systems approach to motor pattern generation. Nature 417:343–350. Ocorr, K.A., and A. Berlind. 1983. The identification and localization of a catecholamine in the motor neurons of the lobster cardiac ganglion. Journal of Neurobiology 14:51–59. Oginsky, M.F., E.W. Rodgers, M.C. Clark, R. Simmons, W.D. Krenz, and D.J. Baro. 2010. D2 receptors receive paracrine neurotransmission and are consistently targeted to a subset of synaptic structures in an identified neuron of the crustacean stomatogastric nervous system. Journal of Comparative Neurology 518:255–276. Pasztor, V.M. 1989. Modulation of sensitivity in invertebrate sensory receptors. Seminars in the Neurosciences 1:5–14. Pedetta, S., L. Kaczer, and H. Maldonado. 2010. Individual aggressiveness in the crab Chasmagnathus: influence in fight outcome and modulation by serotonin and octopamine. Physiology and Behavior 101:438–445. Prinz, A.A., D. Bucher, and E. Marder. 2004. Similar network activity from disparate circuit parameters. Nature Neuroscience 7:1345–1352. Reyes-Colón, D., N. Vázquez-Acevedo, N.M. Rivera, S.H. Jezzini, J. Rosenthal, E.A. Ruiz-Rodríguez, D.J. Baro, A.B. Kohn, L.L. Moroz, and M.A. Sosa. 2010. Cloning and distribution of a putative octopamine/tyramine receptor in the central nervous system of the freshwater prawn Macrobrachium rosenbergii. Brain Research 1348:42–54. Russell, D.F., and D.K. Hartline. 1978. Bursting neural networks: a reexamination. Science 200:453–456. Saver, M.A., J.L. Wilkens, and N.I. Syed. 1999. In situ and in vitro identification and characterization of cardiac ganglion neurons in the crab, Carcinus maenas. Journal of Neurophysiology 81:2964–2976. Scholz, N.L., J. De Vente, J.W. Truman, and K. Graubard. 2001. Neural network partitioning by NO and cGMP. Journal of Neuroscience 21:1610–1618. Selcho, M., D. Pauls, B.E. Jundi, R.F. Stocker, and A.S. Thum. 2012. The role of octopamine and tyramine in Drosophila larval locomotion. Journal of Comparative Neurology 520:3764–3785. Selverston, A.I., D.F. Russell, J.P. Miller, and D.G. King. 1976. The stomatogastric nervous system: structure and function of a small neural network. Progress in Neurobiology 7:215–290. Skiebe, P. 1999. Allatostatin-like immunoreactivity in the stomatogastric nervous system and the pericardial organs of the crab Cancer pagurus, the lobster Homarus americanus, and the crayfish Cherax destructor and Procambarus clarkii. Journal of Comparative Neurology 403:85–105. Skiebe, P., and H. Schneider. 1994. Allatostatin peptides in the crab stomatogastric nervous system: inhibition of the pyloric motor pattern and distribution of allatostatin-like immunoreactivity. Journal of Experimental Biology 194:195–208.



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Skiebe, P., and T. Wollenschläger. 2002. Putative neurohemal release zones in the stomatogastric nervous system of decapod crustaceans. Journal of Comparative Neurology 453:280–291. Smarandache, C.R., and W. Stein. 2007. Sensory-induced modification of two motor patterns in the crab, Cancer pagurus. Journal of Experimental Biology 210:2912–2922. Sosa, M.A., N. Spitzer, D.H. Edwards, and D.J. Baro. 2004. A crustacean serotonin receptor: cloning and distribution in the thoracic ganglia of crayfish and freshwater prawn. Journal of Comparative Neurology 473:526–537. Spitzer, N., D.H. Edwards, and D.J. Baro. 2008. Conservation of structure, signaling and pharmacology between two serotonin receptor subtypes from decapod crustaceans, Panulirus interruptus and Procambarus clarkii. Journal of Experimental Biology 211:92–105. Stein, W. 2009. Modulation of stomatogastric rhythms. Journal of Comparative Physiology A 195:989–1009. Stein, W., N.D. Delong, D.E. Wood, and M.P. Nusbaum. 2007. Divergent co-transmitter actions underlie motor pattern activation by a modulatory projection neuron. European Journal of Neuroscience 26:1148–1165. Stuart, A.E., J. Borycz, and I.A. Meinertzhagen. 2007. The dynamics of signaling at the histaminergic photoreceptor synapse of arthropods. Progress in Neurobiology 82:202–227. Sullivan, R.E., and M.W. Miller. 1984. Dual effects of proctolin on the rhythmic burst activity of the cardiac ganglion. Journal of Neurobiology 15:173–196. Sullivan, R.E., and M.W. Miller. 1990. Cholinergic activation of the lobster cardiac ganglion. Journal of Neurobiology 21:639–650. Swensen, A.M., and E. Marder. 2000. Multiple peptides converge to activate the same voltage-dependent current in a central pattern-generating circuit. Journal of Neuroscience 20:6752–6759. Swensen, A.M., and E. Marder. 2001. Modulators with convergent cellular actions elicit distinct circuit outputs. Journal of Neuroscience 21:4050–4058. Szucs, A., H.D. Abarbanel, M.I. Rabinovich, and A.I. Selverston. 2005. Dopamine modulation of spike dynamics in bursting neurons. European Journal of Neuroscience 21:763–772. Thirumalai, V., A.A. Prinz, C.D. Johnson, and E. Marder. 2006. Red pigment concentrating hormone strongly enhances the strength of the feedback to the pyloric rhythm oscillator but has little effect on pyloric rhythm period. Journal of Neurophysiology 95:1762–1770. Thuma, J.B., L.G. Morris, A.L. Weaver, and S.L. Hooper. 2003. Lobster (Panulirus interruptus) pyloric muscles express the motor patterns of three neural networks, only one of which innervates the muscles. Journal of Neuroscience 23:8911–8920. Tschuluun, N., W.M. Hall, and B. Mulloney. 2009. State-changes in the swimmeret system: a neural circuit that drives locomotion. Journal of Experimental Biology 212:3605–3611. Vázquez-Acevedo, N., D. Reyes-Cólon, E.A. Ruiz-Rodríguez, N.M. Rivera, J. Rosenthal, A.B. Kohn, L.L. Moroz, and M.A. Sosa. 2009. Cloning and immunoreactivity of the 5-HT1Mac and 5-HT2Mac receptors in the central nervous system of the freshwater prawn Macrobrachium rosenbergii. Journal of Comparative Neurology 513:399–416. Weimann, J.M., P. Meyrand, and E. Marder. 1991. Neurons that form multiple pattern generators: identification and multiple activity patterns of gastric/pyloric neurons in the crab stomatogastric system. Journal of Neurophysiology 65:111–122. Wilkens, J.L., T. Shinozaki, T. Yazawa, and H.E.D.J. ter Keurs. 2005. Sites and modes of action of proctolin and the FLP F2 on lobster cardiac muscle. Journal of Experimental Biology 208:737–747. Wood, D.E., and M.P. Nusbaum. 2002. Extracellular peptidase activity tunes motor pattern modulation. Journal of Neuroscience 22:4185–4195. Wood, D.E., W. Stein, and M.P. Nusbaum. 2000. Projection neurons with shared cotransmitters elicit different motor patterns from the same neural circuit. Journal of Neuroscience 20:8943–8953. Wood, D.E., Y. Manor, F. Nadim, and M.P. Nusbaum. 2004. Intercircuit control via rhythmic regulation of projection neuron activity. Journal of Neuroscience 24:7455–7463. Yamagishi, H., H. Miyamoto, and A. Sakurai. 2004. Developmental changes in dopamine modulation of the heart in the isopod crustacean Ligia exotica: reversal of chronotropic effect. Zoological Science 21:917–922.

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Ronald Harris-Warrick Yeh, S.R., R.A. Fricke, and D.H. Edwards. 1996. The effect of social experience on serotonergic modulation of the escape circuit of crayfish. Science 271:366–369. Zhang, B., and R.M. Harris-Warrick. 1994. Multiple receptors mediate the modulatory effects of serotonergic neurons in a small neural network. Journal of Experimental Biology 190:55–77. Zhang, B., and R.M. Harris-Warrick. 1995. Calcium-dependent plateau potentials in a crab stomatogastric ganglion motoneuron. I. Calcium current and its modulation by serotonin. Journal of Neurophysiology 74:1929–1937. Zhang, B., J.F. Wooton, and R.M. Harris-Warrick. 1995. Calcium-dependent plateau potentials in a crab stomatogastric ganglion motoneuron. II. Calcium-activated slow inward current. Journal of Neurophysiology 74:1938–1946. Zhang, H., E.W. Rodgers, W.D. Krenz, M.C. Clark, and D.J. Baro. 2010. Cell specific dopamine modulation of the transient potassium current in the pyloric network by the canonical D1 receptor signal transduction cascade. Journal of Neurophysiology 104:873–884. Zhao, S., J. Golowasch, and F. Nadim. 2010. Pacemaker neuron and network oscillations depend on a neuromodulator-regulated linear current. Frontiers in Behavioral Neuroscience 4:21.

6 SYNAPSES IN CRUSTACEANS

Gregory Lnenicka

Abstract The study of crustacean synapses has a long history and contributed many notable advances to our understanding of synaptic structure and function. Most of these studies were performed in the neuromuscular system due to the large, identifiable motor axons and the accessibility of the neuromuscular synapses. The identification and characterization of glutamate and gammaaminobutyric acid as the excitatory and inhibitory neurotransmitters at neuromuscular synapses played an important role in our understanding of these neurotransmitters and studies in the central nervous system have characterized the action of various other neurotransmitters. The first electrical synapse was described in the crustacean central nervous system, and presynaptic inhibition was originally demonstrated at the neuromuscular synapses; subsequent studies examined the mechanisms of presynaptic inhibition and the functional role of electrical synapses. Crustacean synapses show various forms of activity-dependent synaptic enhancement and depression previously described for vertebrate synapses. In addition the motor terminals show a long-term facilitation, which appears unique to crustaceans. Our knowledge of the role of Ca 2+ in transmitter release particularly during synaptic enhancement was significantly advanced by crustacean studies in which Ca 2+ indicators were injected into motor axons to image Ca 2+ changes in the terminals. Crustacean neuromuscular synapses are well known for their diverse transmitter-releasing properties, either when comparing terminals from a single axon or from different axons; this has led to a number of important studies correlating transmitter release with synaptic ultrastructure. Motor axons with large differences in action potential activity levels develop differences in the structure and function of their neuromuscular synapses. Experimental changes in action potential activity indicate that this synaptic differentiation results from long-term, activity-dependent plasticity.

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INTRODUCTION Crustacean synapses have been extensively studied for decades and have provided a number of seminal contributions to our knowledge of electrical and chemical synaptic transmission. The number of “firsts” in the study of synapses attributed to crustacean studies is quite impressive, and findings at crustacean synapses often foreshadowed similar findings in other organisms by months to years. For example, studies in the opener muscle showing the first direct demonstration of presynaptic inhibition (Dudel and Kuffler 1961a) were followed in the same year by a report of presynaptic inhibition in the mammalian spinal cord (Eccles 1961), whereas, multiple decades separated the observation that a single axon could produce synapses with diverse transmitter-releasing properties in the crustacean neuromuscular system (Hoyle and Wiersma 1958, Atwood 1967) and similar findings in the mammalian brain (Scanziani et al. 1998). The crustacean neuromuscular junction was an early favorite of synaptic physiologists, who were initially attracted by these accessible synapses formed by identified axons on large muscle fibers. The identified axons allowed the “same” synapses to be analyzed physiologically and morphologically, which led to multiple studies correlating structure and function at these synapses. In fact, crustacean neuromuscular synapses are among the best characterized synapses, both physiologically and morphologically. An important strength of this neuromuscular system was the variety of questions that could be addressed, many of them relevant to the central nervous system. The system contains both inhibitory and excitatory synapses, a diversity of transmitter-releasing properties, short and long-term forms of activity-dependent plasticity and regulation by neuromodulators. An additional technical advantage provided by this system was the relatively large size of the axons, which allowed intracellular recording of the presynaptic action potential (AP) and the injections of various inhibitor and activators of synaptic function as well as Ca2+ indicators. A number of techniques were important for advancing our knowledge of crustacean synapses, and this system played an important role in the development of these techniques. One of the most important electrophysiological techniques was extracellular focal recording from synapses, which was used to its greatest advantage in this system. Extracellular focal recording at synapses was first applied to the frog neuromuscular junction (del Castillo and Katz 1956), but after its application to crustacean neuromuscular synapses (Dudel and Kuff ler 1961b) it quickly became a mainstay for quantal studies of synaptic function. This technique allowed an accurate measurement of quantal events, which was not possible using intracellular recording since the innervation is distributed and spontaneous miniature excitatory postsynaptic potentials (EPSPs) could originate length constant(s) apart. A  powerful feature of this approach was that the number of quanta evoked by each AP could be determined from the inf lections on the rising phase of the synaptic currents. Eventually, the advantages of recording transmitter release from just a few boutons were applied to structure-function relationships where the recording sites were marked for subsequent electron microscopy (Wojtowicz et al. 1994). This followed a series of productive ultrastructural studies in which serial-section electron microscopy was used to produce three-dimensional reconstructions of lengths of motor terminals beginning with the crayfish opener muscle (Jahromi and Atwood 1974). Finally, the critical studies using inorganic Ca 2+ indicators to study presynaptic [Ca 2+]i dynamics began with experiments on the excitor motor terminals in the crayfish opener muscle (Delaney et al. 1989). This review of crustacean synapses will focus on neuromuscular synapses although an attempt will be made to relate findings at neuromuscular synapses to central nervous system (CNS) synapses. Studies of synapses in the CNS have provided vital information on the synaptic interactions responsible for reflexes and patterned motor output, as is presented in ­chapters 13–15 in this volume.



Synapses in Crustaceans

SYNAPSES AND NEUROTRANSMITTERS Electrical Synapses Electrical synapses were first discovered in the crustacean nervous system:  electrical transmission was demonstrated at the giant synapse in the crayfish abdomen (Furshpan and Potter 1957) and soon after, electrical synapses were also found connecting large cells in the lobster cardiac ganglion (Watanabe 1958). For the giant synapse, simultaneous intracellular recording from the lateral and motor giant axons in the abdominal nerve cord showed that depolarization, but not hyperpolarization, could be transferred across the synapse from the lateral to motor giant axons (Furshpan and Potter 1959a). This led to the conclusion that the giant synapse is electrical and rectifying, such that current passes only from the pre- to postsynaptic axon. Subsequent studies of the abdominal nerve cord have shown that the synapses formed by mechanosensory afferents and interneurons on the lateral giant are also rectifying electrical synapses (Edwards et  al. 1991). These electrical synapses provide rapid and reliable transmission for the tail-flip escape circuit and also favor the summation of coincident activity for the mechanosensory input to the lateral giant (Edwards et al. 1998). Electrical synapses occur throughout the crustacean CNS. Reciprocal electrical synapses have been demonstrated between primary afferent axons from leg stretch receptors in the crayfish; these synapses may act to synchronize afferent firing to produce greater EPSP summation in postsynaptic cells (El Manira et al. 1993). Motor neurons innervating the same muscle in crayfish walking legs are electrically coupled (Chrachri and Clarac 1989). Also, most neurons in the stomatogastric ganglion have electrical synapses and some have been shown to be rectifying (Nusbaum and Beenhakker 2002). Many of the electrical synapses in the crustacean CNS adjoin chemical synapses to form mixed electrical-chemical synapses; this synaptic configuration may function to allow greater synaptic modulation (Johnson et al. 1994). Excitatory Chemical Synapses Glutamate is the major excitatory neurotransmitter at crustacean neuromuscular synapses. Early studies showed that perfusion of the crayfish claw opener muscle with L-glutamate resulted in contraction (Robbins 1959, van Harreveld and Mendelson 1959). It was subsequently shown that glutamate produced its action at neuromuscular synapses; glutamate was systematically applied by iontophoresis at numerous points along the surface of the crayfish leg opener muscle fiber to reveal sharply localized sites where depolarization occurred (Takeuchi and Takeuchi 1964). By determining the location of the neuromuscular synapses using focal extracellular recording, it was shown that the glutamate-sensitive spots coincided with the neuromuscular synapses (Fig. 6.1). Further evidence for glutamate as the neurotransmitter came from voltage clamp studies of the crayfish claw opener muscle that showed the same reversal potential for the excitatory postsynaptic current (EPSC) and ionophoretically applied glutamate (Onodera and Takeuchi 1975). Finally, it was established that excitatory nerve terminals released glutamate when stimulated. Perfusates collected from crayfish superficial flexor muscles contained more glutamate with nerve stimulation, and less when the external Ca 2+ concentration was lowered to reduce transmitter release (Kawagoe et al. 1981). The postsynaptic action of glutamate was characterized by voltage clamping the muscle fibers; these macroscopic currents showed that glutamate produced an equal increase in Na+ and K+ permeability, which was greater than twice that seen for Ca 2+ and Mg2+ (Dekin 1983). These relative ionic permeabilities are similar to those described for the nicotinic acetylcholine-activated channel found at the vertebrate neuromuscular junction (NMJ). The

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Fig. 6.1. Experiments demonstrating that the glutamate-sensitive sites on crayfish opener muscle fibers occur at neuromuscular synapses. (A) The location of neuromuscular synapses was determined by extracellular focal recording. upper trace: The appearance of a negative potential indicated that the extracellular electrode was positioned over the site of the inward synaptic current. lower trace: Simultaneous intracellular recording of EPSPs from the muscle fiber. (B) upper trace: A glutamate-filled pipette was positioned next to the extracellular recording electrode and a current pulse was passed to eject glutamate. middle trace: Activation of glutamate receptors resulted in an inward synaptic current beneath the extracellular electrode. lower trace: Simultaneous intracellular recording of glutamate-induced synaptic potentials from the muscle. Adapted from Takeuchi and Takeuchi 1964, with permission from The Physiological Society.

neuromuscular glutamate receptors are of the quisqualate type:  sensitive to quisqualic acid, but insensitive to kainic acid and N-methyl-D-aspartic acid (Shinozaki 1980). Single-channel records from excised patches of crayfish muscle showed that the selectivity of single ion channels matched that determined by the macroscopic whole-cell currents (Hatt et al. 1988). These channels produced currents of approximately 8 pA (conductance 100 pS) and could be divided into four types based on their rate of opening and desensitization (Dudel et al. 1990a); however, it is not known whether all the channel types are present at neuromuscular synapses. During the production of the miniature EPSCs, it was estimated that the glutamate concentration at the postsynaptic receptors reached over 1 mM, opening 500 postsynaptic channels. Decay of the miniature EPSC most likely resulted from the decline in glutamate concentration and possibly also from channel desensitization (Dudel et al. 1990b). There is a single exception to glutamate’s role as the excitatory neurotransmitter at crustacean neuromuscular synapses. Acetylcholine acts as the excitatory neurotransmitter for neuromuscular synapses found on the extrinsic muscles of the gastric mill; the pharmacology of these receptors is similar to that of the vertebrate nicotinic acetylcholine receptor (Marder and Paupardin-Tritsch 1980). The intrinsic muscles of the gastric mill are glutamatergic; however many of these muscles also have extrajunctional acetylcholine receptors (Lingle 1980). Both glutamate and acetylcholine can function as excitatory neurotransmitters in the crustacean central nervous system. Acetylcholine is the excitatory neurotransmitter released from sensory afferents in crustaceans (Barker et al. 1972); here, it acts on nicotinic receptors (Miller et al. 1992) or on both nicotinic and muscarinic receptors (Le Bon-Jego et al. 2006). Glutamate acts as an excitatory neurotransmitter in the stomatogastric ganglion; however, its action appears to be mainly through the activation of metabotropic receptors (Levi and Selverston 2006). Both acetylcholine and glutamate can also act as inhibitory neurotransmitters in the CNS.



Synapses in Crustaceans

Inhibitory Chemical Synapses Inhibition at crustacean neuromuscular synapses is produced by gamma-aminobutyric acid (GABA). The first evidence for GABA as the inhibitory neurotransmitter came from studies in which an extract (factor I) from the brain and spinal cord of mammals inhibited the crayfish stretch receptor and muscle (Florey 1954). It was proposed that the active ingredient of factor I was GABA (Bazemore et al. 1956) and, although Florey subsequently rejected this idea (Florey 1991), this proposal resulted in subsequent studies of GABA. Extraction and fractionation techniques were used to identify compounds in the lobster CNS that inhibited neuromuscular excitation in crayfish (Dudel et al. 1963). GABA was the most active compound and it was found at highest concentration in nerve bundles containing the largest proportion of inhibitory fibers (Kravitz et al. 1963b). Subsequent work on isolated axons showed that GABA was found in inhibitory axons, but not found in excitatory ones (Kravitz et al. 1963a). In addition, GABA was shown to inhibit AP activity and increase the membrane conductance of the crayfish stretch receptor (Kuffler and Edwards 1958). The basis for its inhibitory action was clearly demonstrated at the crayfish opener muscle: both stimulation of the inhibitor and GABA application were found to increase the membrane conductance to Cl- as shown by the effects of ion substitution on the inhibitory reversal potential (Boistel and Fatt 1958). The site of action of GABA was localized near the glutamate-sensitive sites along the crayfish opener muscle (Takeuchi and Takeuchi 1965); since the inhibitory and excitatory terminals ran in parallel, this was consistent with GABA acting at inhibitory synapses. In addition to its action at peripheral synapses, GABA is also a common inhibitory neurotransmitter in the crustacean CNS. The first evidence for this came from studies of the motor giant motor neuron in the abdominal nerve cord; in addition to its electrical connections, the motor giant receives inhibitory chemical synapses, which were shown to use GABA as the neurotransmitter (Furshpan and Potter 1959b). Glutamate also acts as an inhibitory neurotransmitter in the crustacean CNS. Iontophoresis of either GABA or glutamate produced inhibitory responses in motor neurons of the crab stomatogastric ganglion (Marder and Paupardin-Tritsch 1978) and crayfish motor neurons innervating the leg (Pearlstein et al. 1994), swimmerets (Sherff and Mulloney 1996) and abdominal flexors (Heitler et al. 2001). The inhibitory action of glutamate is not limited to motor neurons, since interneurons in the terminal abdominal ganglion of the crayfish are inhibited by glutamate application (Nagayama 2005). In most of these cases, the inhibitory action of GABA or glutamate involves an increase in Cl- conductance; however, there is also evidence for both GABA and glutamate producing an increase in K+ conductance (Marder and Paupardin-Tritsch 1978, Nagayama 2005). Single-channel recording from stomatogastric neurons demonstrated distinct ionotropic receptors for glutamate and GABA (Cleland and Selverston 1998). Acetylcholine was also found to function as an inhibitory neurotransmitter in the stomatogastric ganglion. Here, motor neurons that use acetylcholine to excite the muscle can release this same transmitter to produce inhibition at their intraganglionic synapses (Marder and Eisen 1984). Histamine has an inhibitory action on a broad range of crustacean neurons; for example it acts on neurosecretory cells, sensory neurons, motor neurons, and interneurons to produce an increase in Cl- conductance (Claiborne and Selverston 1984, McClintock and Ache 1989, Hashemzadeh-Gargari and Freschi 1992, Cebada and García 2007). In some cases, histamine may be acting as a neuromodulator; however, there is evidence for its role as an inhibitory neurotransmitter in the lobster stomatogastric nervous system (Claiborne and Selverston 1984). In addition to postsynaptic inhibition, crustacean neuromuscular systems show presynaptic inhibition. The existence of two types of inhibition was first demonstrated by Marmont and Wiersma (1938). They isolated the excitor and inhibitor axons to the crayfish claw opener

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Gregory Lnenicka muscle so that they could be stimulated separately and recorded tension and action currents (electromyograms) from the opener muscle (Fig. 6.2A). Two types of inhibition were identified, and for one the effect was greatest when the inhibitor was stimulated immediately before the excitor. This type of inhibition was subsequently confirmed by intracellular recording from the muscle fibers and it was proposed that the inhibitor reduced EPSP amplitude by releasing a neurotransmitter that competed postsynaptically with the excitatory neurotransmitter to produce a “curarine-like action” (Kuffler and Katz 1946, Fatt and Katz 1953). However, extracellular focal recordings from excitatory synapses in the opener muscle showed that this inhibition was presynaptic, leading to a decrease in transmitter release (Fig. 6.2 B) (Dudel and Kuffler 1961a). Thus, the inhibitory axon can act at the excitatory terminals to reduce transmitter release in addition to producing postsynaptic inhibition; the same inhibitory axon has been shown to produce both presynaptic inhibition and postsynaptic inhibition for a number of crustacean muscles. The inhibitor produces presynaptic inhibition by releasing GABA onto the excitatory motor terminals, thereby activating ionotropic receptors and producing an increase in Cl- conductance and hyperpolarization of the membrane potential (Dudel and Kuffler 1961a, Takeuchi and Takeuchi 1966, Kawai and Niwa 1977). This increase in conductance is thought to shunt the inward Na+ current produced by the arriving AP, which would reduce transmitter release by limiting depolarization and Ca 2+ influx at the motor terminals. This is supported by a reduction in the amplitude of the AP recorded from excitatory motor axons near their motor terminals during presynaptic inhibition (Baxter and Bittner 1991). The axoaxonal synapses are strategically positioned to affect the depolarization of the motor terminals; most axoaxonal synapses on the excitor in the spider crab stretcher muscle were located on synaptic varicosities with a few on bottlenecks and the preterminal axon (Atwood et al. 1984). In addition to ionotropic receptors (GABA A), metabotropic GABA B receptors were found on the excitor motor terminals of the crayfish opener muscle (Fischer and Parnas 1996a). It appears that presynaptic inhibition produced by single APs in the inhibitor is mediated by GABA A receptors and presynaptic inhibition produced by AP trains involves both GABA A and GABA B receptors (Fischer and Parnas 1996b). Some presynaptic receptors at neuromuscular synapses act as autoreceptors and provide feedback inhibition of transmitter release. In the lobster stretcher muscle, glutamate inhibits transmitter release from both inhibitory and excitatory motor terminals though the activation of metabotropic receptors linked to G-proteins that produce an increase in K+ conductance (Miwa et al. 1990, 1993). At the excitatory terminals, this appears to provide negative feedback of transmitter release; however, the function of a glutamate-mediated reduction of transmitter release from inhibitory terminals remains unclear. It was proposed that the glutamate-dependent inhibition of transmitter release seen at crayfish excitatory motor terminals resulted from glutamate binding to the presynaptic glutamate transporter, which produces an increase in Cl- conductance (Dudel and Schramm 2003). Glutamate transporters can act as both transporters and ion channels; they have separate transmembrane domains for glutamate transport and Cl- permeation (Ryan and Vandenberg 2005). A number of examples of presynaptic inhibition have been detected at crustacean central synapses. Presynaptic inhibition of crayfish tactile sensory afferents was shown to be associated with primary afferent depolarization (Kennedy et al. 1974). For crayfish proprioceptor afferents, the primary afferent depolarization resulted from the activation of GABA receptors, producing an increase in Cl- conductance and a reduction in AP amplitude due to shunting of the inward current (Cattaert et al. 1992), which in turn leads to a decrease in transmitter release (Cattaert and El Manira 1999). In some cases, the primary afferent depolarization involved the action of both GABA and histamine (El Manira and Clarac 1994). There may also be presynaptic autoreceptors in the crayfish CNS since muscarinic agonists produced a long-lasting reduction in

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Fig. 6.2. Evidence for presynaptic inhibition in the crayfish opener muscle. (A) This early study provided evidence that the crayfish opener muscle received two types of inhibition and the timing of stimulation was critical for one type of inhibition. top: Experimental setup for stimulation of either the excitor or inhibitor, and recording the opener muscle active currents (electromyograms) and tension. The thin and thick nerve bundles containing the excitor and inhibitor axons, respectively, were exposed in the meropodite, and each bundle was contacted by a pair of electrodes for electrical stimulation (S). An extracellular recording electrode was positioned next to the opener muscle and connected to the amplifier (A) for recording electromyograms. A thread connected the dactyl to an auxotonic lever (L) equipped with a mirror (M) for measuring opener muscle contractions. bottom: Recording of electromyograms during stimulation of the excitor and inhibitor axons. The upper trace shows electromyograms recorded from the opener muscle. In the middle trace, the bars represent the interval from stimulation of the excitor axon to the appearance of the electromyogram and the vertical lines represent stimulation of the inhibitor axon. Stimulation of the inhibitor always produced a reduction in muscle contraction; however, when it occurred immediately before stimulation of the excitor (arrow) it also produced a large decrease in the amplitude of the electromyogram (from Marmont and Wiersma 1938, with permission from The Physiological Society). (B) Evidence for presynaptic inhibition provided by simultaneous intracellular (upper trace) and extracellular (lower trace) recordings from the crayfish opener muscle. 1. Stimulation of the excitatory axon produced an EPSP in the opener muscle; the negative deflections in the extracellular focal recording are produced by the synapic current entering the muscle. The small nerve terminal potential (arrow) in the lower trace is produced by the AP in the excitatory motor terminal. 2. Stimulation of the inhibitory axon before the excitatory axon (note that the inhibitory nerve terminal potential preceded the excitatory one by 1.5 msec) resulted in a decrease in the EPSP amplitude. More importantly, the focal recordings showed more failures of transmission demonstrating that the inhibition was acting presynaptically to reduce transmitter release (from Dudel and Kuffler 1961a, with permission from The Physiological Society).

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Gregory Lnenicka EPSP amplitude at sensory afferent to interneuron cholinergic synapses and no apparent direct effect on the interneuron (Miller et al. 1992).

SYNAPTIC STRUCTURE Neuromuscular Synapses At crustacean neuromuscular synapses, the motor terminal forms numerous branches along the muscle fiber providing “multiterminal innervation.” The motor terminals penetrate muscle invaginations and typically, are completely surrounded by the muscle fiber. Most terminals are varicose and these varicosities or “synaptic boutons” are the site for the majority of synaptic contacts. The synaptic boutons contain a high density of synaptic vesicles and mitochondria and have specialized synaptic regions where the synaptic cleft has a separation of 200 Å (Fig. 6.3A). The “active zones” of transmitter release at these synapses are seen as presynaptic dense bodies with a clustering of synaptic vesicles (Fig. 6.3B); morphological structures representing exocytotic vesicles have been observed at these active zones (Jahromi and Atwood 1974, Pearce et al. 1986). Freeze-fracture electron microscopy of lobster neuromuscular synapses showed that the presynaptic membrane at the active zone contained large intramembranous particles, some of which must represent Ca2+ channels (Pearce et al. 1986). The postsynaptic membrane contains plaques of large intramembranous particles that presumably include the glutamate receptors. The only difference in the structure of inhibitory and excitatory synapse is the arrangement of the intramembranous particles at the postsynaptic membrane (Franzini-Armstrong 1976) and the size and shape of the synaptic vesicles (Fig. 6.3B); that is, inhibitory vesicles are smaller and under some fixation conditions, more irregular in shape than excitatory vesicles (Uchizono 1967, Tisdale and Nakajima 1976). Central Nervous System Synapses In the CNS, the general features of the chemical synapses resemble those at neuromuscular synapses, synaptic vesicles are clustered at dense bodies and there is an accumulation of particles in the presynaptic membrane at the active zone (Fig. 6.3D). Electrical synapses in the CNS have closely apposed pre- and postsynaptic membranes and regular spacing of gap junctions connecting the two membranes (Fig. 6.3C). In freeze-fracture, these gap junctions constitute a hexagonal array (Fig. 6.3D). In crustaceans, vesicles are often found at electrical synapses; these gap-junction vesicles have been shown to be tethered to both sides of the gap junctions and may be involved in signal transduction (Ohta et al. 2011). Vertebrate gap junction channels are composed of connexin proteins; however, invertebrates form their gap junctions from innexins, which have no sequence homology with connexins (Phelan 2005). Two innexins have been cloned in the lobster, and most stomatogastric ganglionic neurons express one or both of these proteins (Ducret et al. 2006).

PRESYNAPTIC CA 2+ REGULATION AND TRANSMITTER RELEASE Intracellular Ca2+ Regulation The role of intracellular Ca 2+ in transmitter release and synaptic plasticity has been extensively studied at crustacean neuromuscular synapses. Current models of transmitter release begin with presynaptic Ca 2+ entry through high-voltage-activated Ca 2+ channels at the active

A

D

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Fig. 6.3. Electron micrographs of chemical and electrical synapses. (A) In the crayfish opener muscle, the excitatory terminal, E, shows a synapse (between white arrows) with densely stained pre- and postsynaptic membranes and the underlying granular sarcoplasm, G. (B) An inhibitory terminal, I, in the crayfish opener muscle with an active zone, which appears as a presynaptic dense body, D. The synaptic vesicles in the inhibitory terminal are more irregular in shape than those in the excitatory terminal (from Jahromi and Atwood 1974, with permission from The Rockefeller University Press). Scale bar for A,B: 250 nm. (C) An electrical synapse between lateral giant axons in the crayfish abdominal nerve cord. The electrical synapse consists of gap junctions formed by innexons in a regular array spanning the pre- and postsynaptic membranes. Characteristically, both sides of the gap junction were lined with vesicles. Scale bar: 100 nm (from Ohta et al. 2011, with permission from Elsevier Press). (D) Freeze fracture of the mixed chemical and electrical synapse between the medial giant and motor giant axons in the crayfish abdominal nerve cord. The inner leaflet of the presynaptic membrane (PF face) at the chemical synapse shows elongated active zones (az) with their intramembranous particles and occasional vesicle openings (arrows). A fracture plan passing through the postsynaptic membrane shows the outer leaflet (EF face) at the adjacent electrical synapse and an array of gap junction particles (gj), presumably innexons. The fracture plane then passes through the presynaptic cytoplasm showing the synaptic vesicles (sv). Scale bar: 200 nm. From Bosch 1990, with permission from John Wiley and Sons.

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Gregory Lnenicka zone to produce a large and local increase in [Ca 2+]i; these Ca 2+ domains trigger transmitter release. Various subtypes of high-voltage-activated Ca 2+ channels have been identified based on their physiological properties and their sensitivity to organic Ca 2+ channel antagonists. The P/Q-type and N-type Ca 2+ channels appear to play a prominent role in transmitter release in vertebrates (Fisher and Bourque 2001), and this also appears to be the case for crustacean neuromuscular synapses. The P/Q-type Ca 2+ channel blocker, ω-Aga-TX-IVA, eliminated transmitter release from the excitatory and inhibitory terminals on the crayfish opener muscle; however, application of the N-type Ca 2+ channel blocker ω-conotoxin had no effect on transmitter release (Araque et al. 1994). The involvement of N-type Ca 2+ channels in transmitter release may be motor-neuron specific since the N-type blocker ω-conotoxin reduced transmitter release from the fast closer excitor but not the slow closer excitor in the crab (Rathmayer et al. 2002). The P-type Ca 2+ channels are found not only at motor terminals but also in other regions of the motor neuron; for example, there is physiological and morphological evidence for P-type channels on the somata of motor neurons (Hong and Lnenicka 1997, French et al. 2002). Endogenous Ca 2+ buffers strongly influence the amplitude and duration of the Ca 2+ signals. Measurements of the Ca 2+-binding ratio (Ca 2+ bound/Ca 2+ remaining free) have shown that usually less than 5% of the Ca 2+ entering during a single AP remains free, the rest is rapidly bound by endogenous fast Ca 2+ buffers (Neher 1995). An estimate of the Ca 2+-binding ratio for crayfish motor terminals gave a value of 600 (Tank et al. 1995); this value is high compared to other terminals; for example, it is 77 for Drosophila larval motor terminals (He and Lnenicka 2011). This high value for the crayfish is likely due to the inclusion of both fast and slow Ca 2+ buffers in the measurement; most measurements of the Ca 2+-binding ratio have considered only fast Ca 2+ buffering. Indeed, it has been proposed that for crayfish motor terminals the decay of Ca 2+ transients produced by single APs is due largely to slow Ca 2+ buffering, and slow and fast Ca 2+ buffers coexist at these terminals (Lin et al. 2005); however, none of the Ca 2+ buffers found at these crustacean terminals have been identified. During trains of APs, Ca 2+ buffers saturate and Ca 2+ clearance results from the uptake of Ca 2+ by mitochondria and Ca 2+ extrusion via the plasma membrane Ca 2+ ATPase and Na+/Ca 2+ exchanger (Mulkey and Zucker 1992, Zhong et al. 2001). Transmitter Release Theoretical studies based on measurements from the squid giant synapse and crayfish neuromuscular synapses predicted that a very high [Ca 2+]i (>100 µM) at the Ca 2+ domains situated at active zones triggered transmitter release (Yamada and Zucker 1992). This was supported by studies using flash photolysis of caged Ca 2+ in retinal bipolar cell terminals, which found that transmitter release required a large increase in [Ca 2+]i (Heidelberger et al. 1994). However, more recent studies of mammalian CNS synapses have shown that [Ca 2+]i increases as low as 1–2 µM can trigger transmitter release; this has led to models where single APs produce brief [Ca 2+]i increases of 10–25 µM to evoke transmitter release (Schneggenburger and Neher 2005). The same may be true at crustacean neuromuscular synapses since photolysis of caged Ca 2+ injected into the presynaptic terminal found that [Ca 2+]i elevations of about 0.5 µM could produce transmitter release (Millar et al. 2005). Crustacean presynaptic terminals have not been well characterized molecularly; however, it is likely that the synaptic proteins involved in transmitter release are similar to those found in mammalian species and in Drosophila. Antibodies against the Drosophila synaptic proteins synaptotagmin, synapsin, dynamin, and frequenin apparently recognize the homologous proteins in crustaceans (Cooper et al. 1995a, Jeromin et al. 1999). Frequenin has been cloned in crustaceans and its amino acid sequence is very similar to that of Drosophila frequenin and 64–70%



Synapses in Crustaceans

similar to rat and Xenopus frequenin sequences (Jeromin et al. 1999). In addition, transmitter release is influenced by the injection of mammalian α-SNAP or antibodies raised against mammalian synaptotagmin into crayfish motor terminals (He et al. 1999, Hua et al. 2007).

ACTIVITY-DEPENDENT SYNAPTIC PLASTICITY Synaptic Enhancement The most common forms of synaptic plasticity are the short-term synaptic enhancements, which appear to be present to some degree at all synapses (Zucker and Regehr 2002). Synaptic enhancement can be divided into various components based on differences in duration and was first described at the frog neuromuscular junction. Synaptic facilitation is the shortest component. It can be measured with paired pulses and is often referred to as paired-pulse facilitation. At many synapses it can be clearly divided into F1 and F2 components; the F1 component has a decay time constant of tens of milliseconds and the F2 component develops more slowly and decays with a time constant of hundreds of milliseconds (Mallart and Martin 1967). The enhancement seen during trains of APs results from the buildup of facilitation plus the onset of the longer-lasting augmentation and posttetanic potentiation (PTP). Augmentation decays with a time constant of 5–10 sec and PTP has a decay time constant of 30 sec to several minutes (Magleby 1973, Magleby and Zengel 1976). The synaptic enhancements described at vertebrate neuromuscular junctions have been demonstrated at many other synapses, including those of crustaceans. Crustacean neuromuscular synapses show both F1 and F2 components of short-term synaptic facilitation as well as augmentation and PTP (Zucker 1974, Delaney et al. 1989, Bittner and Baxter 1991). At crustacean NMJs, these components of synaptic enhancement are attributable to the residual Ca 2+ remaining in the terminal after transmitter release. The best evidence for this comes from studies in which exogenous Ca 2+ buffers were used to reduce residual intracellular Ca 2+ resulting in a reduction in synaptic enhancement (Delaney et  al. 1991, Hochner et  al. 1991, Kamiya and Zucker 1994). The mechanism for synaptic enhancement remains unknown, but it is unlikely to result from the summation of entering Ca 2+ with remaining residual Ca 2+ since residual Ca 2+ is usually less than a micromolar after it equilibrates in the terminal. Probably the best supported model for synaptic facilitation at crustacean neuromuscular synapses is a two-site hypothesis that proposes synaptic facilitation is induced by Ca 2+ binding to a high-affinity facilitation site that is separate from the low-affinity sensor for exocytosis (Matveev et al. 2002). A recent modification of this two-site model proposes that the facilitation site involves the priming of docked vesicles by residual Ca 2+ (Millar et al. 2005, Pan and Zucker 2009). An attractive and simpler hypothesis proposes that synaptic facilitation results from the saturation of fast Ca 2+ buffers, which leads to a greater increase in [Ca 2+]i at Ca 2+ domains during subsequent AP activity. This mechanism is supported by findings at mammalian central synapses (Blatow et al. 2003); however, at this point the model has not been supported by findings at crustacean neuromuscular synapses (Matveev et al. 2004). Long-term facilitation (LTF) is another activity-dependent increase in transmitter release found at crustacean neuromuscular synapses; it is produced by trains of APs delivered at moderate stimulation rates (Sherman and Atwood 1971). The early phase of LTF appears to represent PTP; however LTF also has a persistent phase that lasts for hours (Wojtowicz and Atwood 1985)  and possibly days (Lnenicka and Atwood 1985b). This persistent LTF (subsequently referred to as simply LTF) is homosynaptic and has not been described at other synapses. (The LTF described at Aplysia central synapses is heterosynaptic and presumably acts by a different

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Gregory Lnenicka mechanism.) Posttetanic potentiation largely results from Na+ loading of the terminals (Atwood et al. 1975), which appears to increase presynaptic [Ca 2+]i by reversing the Na/Ca exchanger to produce Ca 2+ influx (Zhong et  al. 2001). The decay of PTP follows the decay in presynaptic [Ca 2+]i; however, LTF continues after [Ca 2+]i has returned to resting levels (Delaney et al. 1989). It remains unclear whether an increase in presynaptic [Ca 2+]i is required for the induction of LTF. Long-term facilitation was not blocked by procedures designed to prevent an increase in presynaptic [Ca 2+]i leading to the proposal that LTF is voltage dependent, a unique mechanism for inducing synaptic plasticity (Wojtowicz and Atwood 1988); however, other techniques for preventing a rise in presynaptic [Ca 2+]i did block LTF (Beaumont et al. 2001). Nonetheless, local protein synthesis at the motor terminals is apparently required to produce LTF since protein translation inhibitors blocked the induction of LTF (Beaumont et al. 2001). The expression of LTF appears to involve the activation of silent synapses. The early electron-microscopy studies of crustacean motor terminals and many subsequent studies have shown that there are many more synapses and active zones than the number of transmitter quanta normally released (Jahromi and Atwood 1974). Later statistical analyses of transmitter release at these synapses found that the number of responding units was much less than the number of synapses or active zones and the number of transmitter release sites increased during LTF (Wojtowicz and Atwood 1986); this could be due to more presynaptic dense bodies per synapse (Wojtowicz et al. 1994). Thus, LTF may show similarities to long-term potentiation seen in the mammalian CNS where there is evidence that silent synapses are activated to produce synapse strengthening (Atwood and Wojtowicz 1999). Synaptic Depression Prolonged high-frequency stimulation can result in a reduction in transmitter release. This high-frequency depression is particularly prominent in phasic motor neurons and probably results from a depletion of synaptic vesicles (Bryan and Atwood 1981). In addition, phasic motor neurons often show low-frequency synaptic depression, which occurs at stimulation frequencies as low as 0.1 Hz and is not due to depletion of transmitter (Bruner and Kennedy 1970, Zucker and Bruner 1977). Although low-frequency depression has not been widely reported, it does occur in other organisms, such as rats and Aplysia (Abrahamsson et al. 2005, Doussau et al. 2010). In crustaceans low-frequency depression involves the dephosphorylation of presynaptic proteins and may result from the activation of calcineurin, resulting in dephosphorylation of actin and tubulin and rearrangement of the cytoskeleton (Silverman-Gavrila et  al. 2005, Silverman-Gavrila and Charlton 2009). Short-term synaptic facilitation and depression have also been described at crustacean CNS synapses; however, the extent to which central synapses express this plasticity does not appear to match that seen at neuromuscular synapses. Short-term synaptic plasticity has been examined at central synapses in the stomatogastric ganglion and plays a role in the regulating the rhythmic motor output of this system (Combes et al. 1999, Mamiya and Nadim 2005). Synaptic depression occurring at synapses formed by mechanosensory afferents onto interneurons contributes to the habituation of the tail-flip response in the crayfish (Zucker 1972, Araki and Nagayama 2003). This synaptic depression occurs at very low frequencies of stimulation and so may have features in common with the low-frequency depression seen at neuromuscular synapses.

SYNAPTIC DIVERSITY Crustacean neuromuscular synapses are noted for the diversity of their structure and function. Synaptic diversity is seen when comparing separate motor axons or different terminals from the



Synapses in Crustaceans

same motor axon. Probably the most dramatic differences are seen when comparing “phasic” axons with very low activity levels to “tonic” axons with high activity levels. The differences seen for single motor axons are usually less dramatic but have been clearly demonstrated in a number of cases. Synaptic Differentiation for Single Axons It was noted early that the terminals from a single crustacean motor axon could show differences in transmitter-releasing properties. These synaptic differences can include variation in the size of EPSPs produced by single APs as well as the response of the synapses to repetitive stimulation (Fig. 6.4). In fact, most motor axons show some variation in their neuromuscular synapses especially when comparing different regions of a muscle and it is customary to identify neuromuscular synapses by both their motor axon and their location in the muscle. The axon innervating the opener muscle in the crab walking leg produces facilitating synapses that release small amounts of transmitter for single APs and nonfacilitating synapses with higher transmitter release (Atwood 1967); the former are usually referred to as low output and the latter are high output. A similar pattern of synaptic differentiation was observed for the excitatory axons innervating the proximal accessory flexor muscle of the lobster walking leg (Frank 1973) and the crayfish limb opener muscle (Bittner 1968), where this differentiation has been extensively studied. Synaptic differentiation for individual axons has also been reported for crustacean muscles that receive multiple axons. The superficial flexors are composed of a sheet of about forty muscle fibers no more than two fibers thick, which are innervated by six axons. Each of the six axons form a medial to lateral gradient of EPSP amplitudes: four axons had their largest postsynaptic potentials on the medial edge and two on the lateral edge (Velez and Wyman 1978a). Although there are fewer reports of synaptic differentiation for single axons in the crustacean CNS than for the neuromuscular system, single CNS neurons also produce synapses that show differences in synaptic facilitation or depression (Combes et al. 1999, Mamiya and Nadim 2005). For example, a mechanoreceptor from the lobster gastric mill forms synapses with different degrees of facilitation on two interneurons in the commissural ganglion. Since these two interneurons project to the gastric mill central pattern generator and trigger alternate motor patterns, the motor pattern can be shifted by the frequency of mechanoreceptor firing (Combes et al. 1999). It is likely that this type of synaptic differentiation is a common feature in the CNS; however, it is much more difficult to identify than at the NMJ. The basis for the differences in transmitter release from terminals of the same axon was examined at the NMJ. The high-output and low-output regions of a single motor terminal were compared ultrastructurally, and it was found that the active zone area per synapse was greater for high-output synapses than low-output ones (Govind and Chiang 1979, Govind and Meiss 1979). However, it was concluded that there was not a linear relationship between active zone area and transmitter release and other factors were likely involved in the differentiation of transmitter release (Atwood and Marin 1983). Freeze-fracture studies of the crayfish opener muscle showed that the active zones for high-output and low-output terminals did not show differences in the number of intramembranous particles, which presumably included Ca 2+ channels (Govind et al. 1994). Nonetheless, Ca 2+-imaging experiments indicated greater Ca 2+ influx at the high-output boutons compared to the low-output ones, and this greater Ca 2+ influx per active zone at high-output boutons could be responsible for their greater transmitter release (Cooper et al. 1995b). It may be that the comparison of intramembranous particles did not reflect the number of Ca 2+ channels or there were differences in the percentage of Ca 2+ channels that were functional at high and low-output boutons. In addition, it has been proposed that the higher output synapses have more closely spaced active zones whose Ca 2+ domains sum to produce a greater Ca 2+ signal and transmitter release (Cooper et al. 1996).

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Fig. 6.4. Variation in synaptic properties for a single axon innervating the stretcher muscle in the walking leg of the spider crab. The EPSPs recorded from three separate fibers show differences in amplitude during stimulation at 1 Hz (A 1, B1, and C1) and when stimulated at 1 Hz, then 10 Hz, and back to 1 Hz (A 2, B2, and C2). The synaptic properties range from low-output, facilitating (A), to high-output, nonfacilitating (C). Calibration: 7.5 mV in A 1 and A 2 and 15 mV in the remaining traces; 30 msec in A 1 and B1, 75 msec in C1, and 750 msec in A 2, B2, and C2 (from Sherman and Atwood 1972, with permission from The Rockfeller University Press).

The differences in synaptic facilitation found at terminals from the same axon could result from their differences in basal transmitter release: high-output synapses would show less facilitation than low-output ones because they are approaching saturation of transmitter release. This was supported by studies of crab neuromuscular synapses, where an experimental increase in basal transmitter release resulted in a decrease in synaptic facilitation (Atwood and Lang 1973). In fact, the differences in facilitation for synapses from the same motor axon can be can be reduced by altering basal transmitter release. A motor axon that projects to two different muscles in the lobster stomatogastric system has high-output and depressing synapses on one muscle and low-output and facilitating synapses on the other. When transmitter release was reduced with saline containing low Ca 2+ and high Mg2+, both terminals produced synaptic facilitation (Katz et al. 1993). It has been proposed that neuromuscular synaptic diversity results from the specification of synaptic properties by the target muscle fibers. The best evidence for this remains a study of the excitatory axon innervating the lobster accessory flexor muscle (Frank 1973). Here motor terminals that innervated separate fibers had different facilitation properties and those that innervated the same fiber had similar facilitation properties. These findings were obtained by selectively stimulating separate primary axonal branches that converged on the same muscle fiber and comparing their synaptic facilitation. The synaptic facilitation for terminals innervating the same fiber was strongly correlated even though they originated from separate branches. Again, the terminals with the greatest synaptic facilitation also had low-output synapses, and it may be that the muscle fiber is specifying basal transmitter release, which then leads to the differences in synaptic facilitation.



Synapses in Crustaceans

Phasic-Tonic Synaptic Differentiation The differences in the structure and function of phasically and tonically active motor terminals can be quite dramatic. The phasic terminals release large amounts of transmitter and produce sizable EPSPs for single APs. The tonic terminals generally release less transmitter and produce much smaller EPSPs. There is also a striking difference in their response to prolonged repetitive stimulation. The tonic terminals often show facilitation and are noted for their ability to maintain transmitter release during stimulation at moderate frequencies for many minutes, whereas transmitter release from phasic terminals depresses and is often negligible after only a minute or so of stimulation (Fig. 6.5A). Also, low-frequency depression is often seen at phasic terminals but is not found at tonic terminals. These synaptic differences can be seen when comparing phasic and tonic motor axons innervating the same muscle fibers or different muscle fibers. There are dramatic morphological differences in the phasic and tonic motor terminals, and these differences appear to be related to their differences in synaptic depression. Phasic terminals, especially those with very low AP activity, are thin and uniform in diameter with few discernible synaptic boutons (Fig. 6.5A) (Lnenicka et al. 1986, 1991). The phasic terminals have relatively few and small mitochondria that are evenly distributed along the terminals (Fig. 6.5B); synapses are also evenly spaced along the phasic terminals and often have relatively small clusters of synaptic vesicles (Atwood and Jahromi 1978, Lnenicka et al. 1986). Tonic terminals are varicose with large synaptic boutons, and the synapses are clustered in these boutons, which contain numerous, large mitochondria and a large population of synaptic vesicles (Jahromi and Atwood 1974, King et al. 1996). Consistent with the differences in synaptic vesicles, tonic terminals have a higher level of the neurotransmitter glutamate than phasic terminals (Shupliakov et al. 1995). Presumably, a smaller reserve of vesicles and lower rates of vesicle recycling lead to more rapid depletion of vesicles and greater synaptic depression for phasic terminals. Here, the differences in mitochondrial content and energy metabolism likely play an important role, since inhibitors of oxidative phosphorylation result in increased synaptic fatigue (Nguyen and Atwood 1997). The basis for the phasic-tonic differences in transmitter release for single APs has been studied at the crayfish extensor muscle of the first walking leg. Here the phasic terminals release 100 to 1000 times more transmitter than the tonic terminals, and this could not be accounted for by differences in synapses, active zones or Ca 2+ entry (King et al. 1996, Msghina et  al. 1999). Instead, the greater transmitter release from phasic terminals was mainly due to greater Ca 2+ sensitivity of transmitter release for phasic compared to tonic synapses (Millar et  al. 2005). This was demonstrated by producing similar increases in [Ca 2+]i in phasic and tonic terminals using photolysis of caged Ca 2+. For the same increase in [Ca 2+]i the release rates for phasic boutons were up to 100 times greater than tonic boutons (Fig. 6.6). The differences between phasic and tonic terminals are neuron specific and not muscle specific. This is evidenced by the fact that phasic and tonic terminals appear on the same muscle fibers, often running side by side (Lnenicka et al. 1986, King et al. 1996). In addition, phasic terminals maintain their properties when transplanted to a tonic muscle. The phasic axons that normally innervate the fast deep flexors muscle were transplanted to the denervated slow superficial flexor muscles. The regenerated phasic axons showed their normal properties: large EPSPs, more profuse innervation, thin terminals and low mitochondrial volume (Krause et al. 1998).

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Fig. 6.5. Phasic and tonic motor terminals. (A) Phasic and tonic terminals innervating the crayfish closer muscle were injected with horseradish peroxidase to show their differences in structure. The varicose tonic terminals produce EPSPs that facilitate during 20-Hz stimulation and maintain transmitter release for prolonged periods. The thin phasic terminals (arrowheads) produce a large EPSPs that show dramatic synaptic depression during 20-Hz stimulation. Calibration: 10 mV, 10 sec. (from Lnenicka and Morley 2002, with permission from Springer). (B) Phasic and tonic motor terminals innervating the crayfish leg extensor were reconstructed from serial electron micrographs. The reconstructions show that the tonic terminal contains three branched mitochondria (m1, m2, and m3), whereas the phasic terminal contains only a single unbranched mitochondria. The dark shaded areas on the presenting surface represent synapses. Scale bar: 1 µm. From King et al. 1996, with permission from John Wiley and Sons.

Fig. 6.6. Differences in the Ca 2+ sensitivity of transmitter release for phasic and tonic motor terminals. The phasic (A) and tonic (B) terminals innervating the extensor muscle in the crayfish walking legs were filled with the caged Ca 2+ chelator DMNPE-4. Bottom: The relative intensity of the UV flash used to release Ca 2+ from DMNPE-4. Middle: The increase in [Ca 2+]i produced by the release of caged Ca 2+. Top: Transmitter release was monitored by extracellular focal recording from phasic or tonic synaptic boutons. The numbers represent the maximum rate of transmitter release (quanta/sec) during the increase in [Ca 2+]i. A similar increase in [Ca 2+]i in phasic and tonic boutons produced a much greater increase in transmitter release in phasic boutons (from Millar et al. 2005, with permission from The Society for Neuroscience).



Synapses in Crustaceans

DEVELOPMENTAL SYNAPTIC PLASTICITY Growth-Related Synaptic Changes Synapses have long been noted for their capacity to change, however mechanisms also exist to maintain a constant synaptic efficacy (synaptic homeostasis). This homeostatic synaptic plasticity functions to compensate for changes resulting from growth, protein turnover, and long-term changes in AP activity (Turrigiano and Nelson 2004). Initial studies of synaptic homeostasis focused on the maintenance of synaptic strength during growth. The crustacean neuromuscular junction provided a suitable model system for these studies since the muscle fibers undergo a large increase in diameter during postnatal growth. It was shown that the EPSP amplitude produced by single APs was maintained during muscle fiber growth due to an increase in transmitter release from the motor terminals in both the lobster walking leg and the crayfish superficial f lexors (DeRosa and Govind 1978, Lnenicka and Mellon 1983a). In addition, studies in the crayfish showed that there was an increase in the amplitude and duration of the miniature EPSCs during growth; this increase in quantal size also contributed to the maintenance of EPSP amplitude (Lnenicka and Mellon 1983a). In the lobster, the increase in transmitter release was associated with growth of the motor terminal and an increase in the total number of synapses (Pearce et al. 1985). The increase in transmitter release appeared to be a response to muscle growth, since experimentally reducing muscle fiber growth delayed the increase in transmitter release in the crayfish (Lnenicka and Mellon 1983b). Single EPSPs recorded from a lobster stomach muscle also showed a constant amplitude during lobster growth; however, there was a lack of synaptic homeostasis for repetitive synaptic activity. Trains of APs produced greater depolarization in juveniles than in adults due to greater synaptic facilitation and temporal summation (Pulver et al. 2005). Activity-Dependent Synaptic Differentiation The role of electrical activity in determining synaptic structure and function was studied at the NMJs formed by phasic and tonic motor axons. Phasic and tonic motor axons often show large differences in AP activity; for example, the phasic axon innervating the crayfish claw closer muscle fires 1 AP per hour and its tonic counterpart innervating the same muscle fires 6,000 APs per hour (Pahapill et  al. 1985). These differences in activity could play a role in the differentiation of their neuromuscular synapses and this was tested by stimulating phasic axons over a period of days with implanted electrodes. Chronic stimulation of the phasic axon resulted in a partial transformation of the neuromuscular synaptic physiology to a more tonic type (Fig. 6.7A). Physiologically, the terminals reduced their initial output of transmitter but became more resistant to synaptic depression during repetitive stimulation (Lnenicka and Atwood 1985a). Claw immobilization produced the opposite effect: an increase in EPSP amplitude and greater synaptic depression (Pahapill et al. 1985). Both the reduction in initial transmitter release and the increase in fatigue-resistance required protein synthesis and were very long lasting, persisting for weeks (Lnenicka and Atwood 1985a, Nguyen and Atwood 1990). In addition, these physiological changes were more readily produced during development since the changes were greater in young animals compared to old animals (Lnenicka and Atwood 1985a). Finally the generation of PTP was also inf luenced by the previous history of AP activity. Normally PTP is more readily produced at phasic neuromuscular synapses than tonic ones (Pahapill et al. 1987); chronic stimulation of a phasic motor axon diminished

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Gregory Lnenicka the subsequent production of PTP, while a chronic decrease in AP activity enhanced PTP (Pahapill et al. 1986). In addition to the physiological changes, chronic stimulation altered the phasic motor terminal morphology to become more similar to tonic terminals (Fig. 6.7B). The phasic terminals became more varicose and contained more and larger mitochondria; in fact, the percentage of cytoplasm occupied by mitochondria became identical to the tonic terminal. After stimulation, the mitochondria and synapses were concentrated in the newly formed synaptic boutons as seen at tonic terminals (Lnenicka et al. 1986, 1991). This increase in mitochondria and their greater proximity to synapses likely contributed to the greater fatigue resistance of the synapses. In fact, the mitochondria in the stimulated axon became more metabolically active in addition to their increased size and the application of mitochondrial inhibitors reversed the increased fatigue resistance seen during long-term adaptation (Nguyen et al. 1994). There were no morphological changes in synapses or active zones that could contribute to the reduction in initial transmitter release. The synaptic changes produced by chronically stimulating a phasic motor axon were termed long-term adaptation and likely reflect multiple, parallel changes in the synapses. For example the reduction in initial transmitter release and the increase in fatigue resistance occurred by separate mechanisms, since they developed at different rates and had different stimulation requirements (Lnenicka and Atwood 1989, Mercier et al. 1992). The synaptic changes seen during long-term adadptation may be similar to those reported at other synapses. For example, the activity-dependent changes in initial transmitter release are consistent with the homeostatic synaptic plasticity produced by prolonged changes in AP activity in mammalian neurons (Turrigiano and Nelson 2004). This activity-dependent homeostatic plasticity is expressed as an increase in EPSP amplitude in response to a prolonged decrease in AP activity and a decrease in EPSP amplitude resulting from a chronic increase in AP activity. This presumably functions

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Fig. 6.7. The changes in crayfish motor neurons produced by chronically altering their AP activity. These changes were produced by in vivo stimulation of a phasic motor neuron (low AP activity to high AP activity); in some cases, the opposite changes were also produced by chronically reducing electrical activity. (A) Simulated EPSPs showing the relative EPSP amplitudes produced by neuromuscular synapses with low AP activity (controls) and after days of chronic stimulation (high AP activity). There was a decrease in the initial EPSP amplitude produced by single APs after chronic stimulation; however during repetitive stimulation there was less synaptic depression, resulting in a larger final EPSP compared to controls. The EPSP amplitude recovered in controls to express strong PTP but the chronically stimulated synapses showed little PTP. The relative EPSP amplitudes are based on studies performed on the fast closer excitor of the crayfish closer muscle (Lnenicka and Atwood 1985a, Pahapill et al. 1986). (B) Chronic stimulation of the fast closer excitor of the crayfish closer muscle resulted in a change in the morphology of the motor terminals; they became more varicose and there was an increase in the size of the mitochondria (Lnenicka et al. 1986). (C) The changes in Ca 2+ regulation produced by chronically increasing the AP activity of a phasic motor neuron. There was a decrease in voltage-dependent Ca 2+ currents (Hong and Lnenicka 1995) and an increase in Ca 2+ clearance (Fengler and Lnenicka 2002) after chronic stimulation.



Synapses in Crustaceans

to maintain a constant synaptic drive, resulting from the combined effects of the frequency of synaptic activation and synaptic strength. The changes in synaptic depression and PTP produced by chronic changes in AP activity could be considered a form of metaplasticity, the plasticity of synaptic plasticity (Abraham and Bear 1996). This term was used to describe findings at mammalian central synapses where prior stimulation inhibited the subsequent induction of long-term potentiation. Crustacean motor neurons show changes in Ca 2+ regulation as a result of chronic alterations in AP activity; these changes maintain intracellular Ca 2+ homeostasis and may underlie some forms of synaptic plasticity (Fig. 6.7C). For example, increased AP activity and Ca 2+ influx in a phasic motor neuron produces a long-term decrease in Ca 2+ currents through P-type Ca 2+ channels (Hong and Lnenicka 1995, 1997). A reduction in Ca 2+ influx through P-type Ca 2+ channels at the motor terminals could produce the reduction in transmitter release seen during long-term adaptation. This appears to be the case, since the activity-dependent reduction in Ca 2+ currents likely occurs throughout the neuron; Ca 2+ influx limited to the axon and terminals results in a reduction in Ca 2+ currents at the cell body (Hong and Lnenicka 1995). Similarly, the reduction in transmitter release can also be produced by remote Ca 2+ influx: Ca 2+ influx localized to the cell body results in the decrease in transmitter release (Hong and Lnenicka 1993). Also both the decrease in Ca 2+ currents and transmitter release require de novo protein synthesis (Hong and Lnenicka 1993, 1995). Tonically active motor axons show more rapid Ca 2+ clearance than phasically active ones (Lnenicka et  al. 1998, Msghina et  al. 1999)  due to greater activity of the Na+/Ca 2+ exchanger and possibly the plasma membrane Ca 2+ ATPase (Rumpal and Lnenicka 2003). These differences are activity dependent, since silencing a tonic motor axon decreases the rate of Ca 2+ clearance and stimulating a phasic motor axon increases the Ca 2+ clearance rate (Lnenicka et al. 1998, Fengler and Lnenicka 2002). These activity-dependent changes in Ca 2+ clearance may be responsible for the metaplasticity of PTP seen at these synapses. Activity-dependent changes in Ca 2+ influx and Ca 2+ clearance could produce this effect by altering the buildup of residual Ca 2+. The effects of altering AP activity demonstrate that differences in the structure and function of phasic and tonic motor terminals are at least partially specified by AP activity; the extent of this activity-dependent differentiation remains to be determined. The activity-induced transformation of the morphology and thus, possibly the increase in fatigue resistance was limited by a lack of increase in terminal size; that is, the phasic terminal became more varicose but the mean cross-sectional area did not increase (Lnenicka et al. 1986). A more complete transformation might occur if activity were altered early in development when the terminals were first forming. The same may be true for basal transmitter release since stimulation of the phasic axon reduced transmitter release but not to the levels seen for tonic terminals. In addition, the reduction in transmitter release from phasic terminals appeared to result from a decrease in Ca 2+ influx and the phasic-tonic differences in transmitter release include differences in the Ca 2+ sensitivity of transmitter release. It remains to be determined whether AP activity, particularly during early development, can influence the Ca 2+ sensitivity of transmitter release. For tonic axons, there may be a more subtle matching of synaptic properties to AP activity patterns than that seen when comparing phasic and tonic axons. Axons that fire brief high-frequency “bursts” of APs appear to show greater synaptic facilitation than those that fire longer trains of APs at lower frequencies. For example, the larger diameter axons innervating the slow flexors fire bursts of APs and show greater facilitation than the smaller axons, which fire long trains of APs (Velez and Wyman 1978b). This relationship is also seen for lobster claws, where the slow closer excitor (SCE) in the cutter claw fires in bursts and shows greater facilitation than the SCE in the crusher claw, which fires longer AP trains (Lnenicka et al. 1988). For lobsters that develop symmetrical cutter claws due to lack of a substratum, the SCE in both

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Gregory Lnenicka claws fires in bursts and both claws show strong synaptic facilitation. It is not known whether synaptic facilitation differentiates as a result of the differences in the pattern of AP activity but it is possible that long trains of APs lead to development of more rapid Ca 2+ clearance and less facilitation.

FUTURE DIRECTIONS The advantages of the crustacean synapses for cellular studies continue to make this preparation unique and there remain important questions that can be addressed using this system. Many synaptic studies are now performed in organisms that allow genetic manipulations in order to identify proteins important for synaptic function. As these essential molecular components are identified, it may prove advantageous to turn back to crustacean synapses to better characterize them in the context of cellular structure and function and information processing. Here, the ability to inject the motor axons with specific activators or peptide inhibitors that act rapidly on presynaptic proteins and pathways could provide an advantage over the genetic approach, where the molecular changes are long-lasting and can produce developmental effects. Of course, molecular studies of crustacean synapses will require the identification of homologous synaptic proteins in crustaceans, and these studies would greatly benefit from sequencing and annotating crustacean genomes. The basis for synaptic diversity is an important field of synaptic studies, and the crustacean neuromuscular system remains arguably the best system for studying the differentiation of transmitter releasing properties. In particular, the molecular mechanism for variations in the Ca 2+ sensitivity of transmitter release is of great interest, and crustacean phasic and tonic motor axons offer synapses with dramatic differences in Ca 2+ sensitivity. It would not be surprising if these differences were further exploited in future studies. There is considerable interest in understanding silent synapses and their recruitment. The crustacean NMJ provides an accessible population of silent synapses that appear to be recruited during activity-dependent synapse strengthening. It seems likely that this system will prove valuable for testing future hypotheses. Finally, crustacean studies have largely focused on presynaptic mechanisms for regulating synaptic strength, and, given findings at other synapses, more attention could be paid to postsynaptic mechanisms in future studies. An important contribution of crustacean synapses not covered in this chapter, but covered in ­chapter 20 by Johnson et al., is their usefulness as a teaching tool. The relative ease of dissection, the ability to maintain the neuromuscular preparations for hours at room temperature, and the opportunity to record synaptic potentials from large muscle fibers makes this arguably the best preparation for teaching synaptic physiology. Many students are introduced to synaptic physiology through this preparation; this alone ensures the use of crustacean synapses for years to come.

CONCLUSIONS For more than fifty years, crustacean studies have played a critical role in advancing our knowledge of synaptic structure and function. This has included pioneering studies on the identification of excitatory and inhibitory neurotransmitters and their course of action at the postsynaptic membrane. Early crustacean studies were key in the identification and characterization of electrical synaptic transmission, presynaptic inhibition, and neuromodulators. A wealth of ultrastructural studies defined the structure-function relationships at the presynaptic terminal,



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particularly the role of the active zone in transmitter release. Studies of activity-dependent synapse strengthening demonstrated long-term changes in synaptic morphology and supported the activation of silent synapses. New approaches for quantifying Ca 2+ dynamics at the presynaptic terminal allowed a better understanding of Ca 2+ regulation and its relationship to transmitter release and synaptic plasticity. Anyone who worked on this system will forever prize those classic preparations that always worked.

ACKNOWLEDGMENTS I thank Dr. Harold Atwood for his valuable comments on an earlier version of this manuscript. Research support was provided by the National Science Foundation.

REFERENCES Abraham, W.C., and M.F. Bear. 1996. Metaplasticity: the plasticity of synaptic plasticity. Trends in Neurosciences 19:126–130. Abrahamsson, T., B. Gustafsson, and E. Hanse. 2005. Synaptic fatigue at the naive perforant path-dentate granule cell synapse in the rat. Journal of Physiology (London) 569:737–750. Araki, M., and T. Nagayama. 2003. Direct chemically mediated synaptic transmission from mechanosensory afferents contributes to habituation of crayfish lateral giant escape reaction. Journal of Comparative Physiology A 189:731–739. Araque, A., F. Clarac, and W. Buno. 1994. P-type Ca2+ channels mediate excitatory and inhibitory synaptic transmitter release in crayfish muscle. Proceedings of the National Academy of Sciences of the United States of America 91:4224–4228. Atwood, H.L. 1967. Variation in physiological properties of crustacean motor synapses. Nature 215:57–58. Atwood, H.L., and S.S. Jahromi. 1978. Fast-axon synapses of a crab leg muscle. Journal of Neurobiology 9:1–15. Atwood, H.L., and F. Lang. 1973. Differential responses of crab neuromuscular synapses to cesium ion. Journal of General Physiology 61:747–766. Atwood, H.L., and L. Marin. 1983. Ultrastructure of synapses with different transmitter-releasing characteristics on motor axon terminals of a crab, Hyas areneas. Cell and Tissue Research 231:103–115. Atwood, H.L., and J.M. Wojtowicz. 1999. Silent synapses in neural plasticity: current evidence. Learning and Memory 6:542–571. Atwood, H.L., L.E. Swenarchuk, and C.R. Gruenwald. 1975. Long-term synaptic facilitation during sodium accumulation in nerve terminals. Brain Research 100:198–202. Atwood, H.L., J.K. Stevens, and L. Marin. 1984. Axoaxonal synapse location and consequences for presynaptic inhibition in crustacean motor axon terminals. Journal of Comparative Neurology 225:64–74. Barker, D.L., E. Herbert, J.G. Hildebrand, and E.A. Kravitz. 1972. Acetylcholine and lobster sensory neurones. Journal of Physiology (London) 226:205–229. Baxter, D.A., and G.D. Bittner. 1991. Synaptic plasticity at crayfish neuromuscular junctions: presynaptic inhibition. Synapse 7:244–251. Bazemore, A., K.A. Elliott, and E. Florey. 1956. Factor I and gamma-aminobutyric acid. Nature 178:1052–1053. Beaumont, V., N. Zhong, R. Fletcher, R.C. Froemke, and R.S. Zucker. 2001. Phosphorylation and local presynaptic protein synthesis in calcium- and calcineurin-dependent induction of crayfish long-term facilitation. Neuron 32:489–501. Bittner, G.D. 1968. Differentiation of nerve terminals in the crayfish opener muscle and its functional significance. Journal of General Physiology 51:731–758.

167

168

Gregory Lnenicka Bittner, G.D., and D.A. Baxter. 1991. Synaptic plasticity at crayfish neuromuscular junctions: facilitation and augmentation. Synapse 7:235–243. Blatow, M., A. Caputi, N. Burnashev, H. Monyer, and A. Rozov. 2003. Ca2+ buffer saturation underlies paired pulse facilitation in calbindin-D28k-containing terminals. Neuron 38:79–88. Boistel, J., and P. Fatt. 1958. Membrane permeability change during inhibitory transmitter action in crustacean muscle. Journal of Physiology (London) 144:176–191. Bosch, E. 1990. Ultrastructure of the electrotonic and chemical components of the lateral-to-motor and medial-to-motor synapses in crayfish nerve cord. Journal of Comparative Neurology 299:446–461. Bruner, J., and D. Kennedy. 1970. Habituation: occurrence at a neuromuscular junction. Science 169:92–94. Bryan, J.S., and H.L. Atwood. 1981. Two types of synaptic depression at synapses of a single crustacean motor axon. Marine Behaviour and Physiology 8:99–121. Cattaert, D., and A. El Manira. 1999. Shunting versus inactivation: analysis of presynaptic inhibitory mechanisms in primary afferents of the crayfish. Journal of Neuroscience 19:6079–6089. Cattaert, D., A. El Manira, and F. Clarac. 1992. Direct evidence for presynaptic inhibitory mechanisms in crayfish sensory afferents. Journal of Neurophysiology 67:610–624. Cebada, J., and U. García. 2007. Histamine operates Cl--gated channels in crayfish neurosecretory cells. Journal of Experimental Biology 210:3962–3969. Chrachri, A., and F. Clarac. 1989. Synaptic connections between motor neurons and interneurons in the fourth thoracic ganglion of the crayfish, Procambarus clarkii. Journal of Neurophysiology 62:1237–1250. Claiborne, B.J., and A.I. Selverston. 1984. Histamine as a neurotransmitter in the stomatogastric nervous system of the spiny lobster. Journal of Neuroscience 4:708–721. Cleland, T.A., and A.I. Selverston. 1998. Inhibitory glutamate receptor channels in cultured lobster stomatogastric neurons. Journal of Neurophysiology 79:3189–3196. Combes, D., P. Meyrand, and J. Simmers. 1999. Dynamic restructuring of a rhythmic motor program by a single mechanoreceptor neuron in lobster. Journal of Neuroscience 19:3620–3628. Cooper, R.L., D.R. Hampson, and H.L. Atwood. 1995a. Synaptotagmin-like expression in the motor nerve terminals of crayfish. Brain Research 703:214–216. Cooper, R.L., L. Marin, and H.L. Atwood. 1995b. Synaptic differentiation of a single motor neuron: conjoint definition of transmitter release, presynaptic calcium signals, and ultrastructure. Journal of Neuroscience 15:4209–4222. Cooper, R.L., J.L. Winslow, C.K. Govind, and H.L. Atwood. 1996. Synaptic structural complexity as a factor enhancing probability of calcium-mediated transmitter release. Journal of Neurophysiology 75:2451–2466. Dekin, M.S. 1983. Permeability changes induced by L-glutamate at the crayfish neuromuscular junction. Journal of Physiology (London) 341:105–125. del Castillo, J., and B. Katz. 1956. Localization of active spots within the neuromuscular junction of the frog. Journal of Physiology (London) 132:630–649. Delaney, K.R., R.S. Zucker, and D.W. Tank. 1989. Calcium in motor nerve terminals associated with posttetanic potentiation. Journal of Neuroscience 9:3558–3567. Delaney, K., D.W. Tank, and R.S. Zucker. 1991. Presynaptic calcium and serotonin-mediated enhancement of transmitter release at crayfish neuromuscular junction. Journal of Neuroscience 11:2631–2643. DeRosa, R.A., and C.K. Govind. 1978. Transmitter output increases in an identifiable lobster motoneurone with growth of its muscle fibres. Nature 273:676–678. Doussau, F., Y. Humeau, F. Benfenati, and B. Poulain. 2010. A novel form of presynaptic plasticity based on the fast reactivation of release sites switched off during low-frequency depression. Journal of Neuroscience 30:16679–16691. Ducret, E., H. Alexopoulos, F.Y. Le, J.A. Davies, P. Meyrand, J.P. Bacon, and V.S. Fenelon. 2006. Innexins in the lobster stomatogastric nervous system: cloning, phylogenetic analysis, developmental changes and expression within adult identified dye and electrically coupled neurons. European Journal of Neuroscience 24:3119–3133.



Synapses in Crustaceans

Dudel, J., and S.W. Kuffler. 1961a. Presynaptic inhibition at the crayfish neuromuscular junction. Journal of Physiology (London) 155:543–562. Dudel, J., and S.W. Kuffler. 1961b. The quantal nature of transmission and spontaneous miniature potentials at the crayfish neuromuscular junction. Journal of Physiology (London) 155:514–529. Dudel, J., and M. Schramm. 2003. A receptor for presynaptic glutamatergic autoinhibition is a glutamate transporter. European Journal of Neuroscience 18:902–910. Dudel, J., R. Gryder, A. Kaji, S.W. Kuffler, and D.D. Potter. 1963. Gamma-aminobutyric acid and other blocking compounds in Crustacea. I. Central nervous system. Journal of Neurophysiology 26:721–728. Dudel, J., C.A. Frank, and H. Hatt. 1990a. A family of glutamergic, excitatory channel types at the crayfish neuromuscular junction. Journal of Comparative Physiology A 166:757–768. Dudel, J., C. Franke, and H. Hatt. 1990b. Rapid activation, desensitization, and resensitization of synaptic channels of crayfish muscle after glutamate pulses. Biophysical Journal 57:533–545. Eccles, J.C. 1961. The Ferrier lecture: The nature of central inhibition. Proceedings of the Royal Society of London B: Biological Sciences 153:445–476. Edwards, D.H., W.J. Heitler, E.M. Leise, and R.A. Fricke. 1991. Postsynaptic modulation of rectifying electrical synaptic inputs to the LG escape command neuron in crayfish. Journal of Neuroscience 11:2117–2129. Edwards, D.H., S.R. Yeh, and F.B. Krasne. 1998. Neuronal coincidence detection by voltage-sensitive electrical synapses. Proceedings of the National Academy of Sciences of the United States of America 95:7145–7150. El Manira, A., and F. Clarac. 1994. Presynaptic inhibition is mediated by histamine and GABA in the crustacean escape reaction. Journal of Neurophysiology 71:1088–1095. El Manira, A., D. Cattaert, P. Wallen, R.A. DiCaprio, and F. Clarac. 1993. Electrical coupling of mechanoreceptor afferents in the crayfish: a possible mechanism for enhancement of sensory signal transmission. Journal of Neurophysiology 69:2248–2251. Fatt, P., and B. Katz. 1953. The effect of inhibitory nerve impulses on a crustacean muscle fibre. Journal of Physiology (London) 121:374–389. Fengler, B.T., and G.A. Lnenicka. 2002. Activity-dependent plasticity of calcium clearance from crayfish motor axons. Journal of Neurophysiology 87:1625–1628. Fischer, Y., and I. Parnas. 1996a. Activation of GABAB receptors at individual release boutons of the crayfish opener neuromuscular junction produces presynaptic inhibition. Journal of Neurophysiology 75:1377–1385. Fischer, Y., and I. Parnas. 1996b. Differential activation of two distinct mechanisms for presynaptic inhibition by a single inhibitory axon. Journal of Neurophysiology 76:3807–3816. Fisher, T.E., and C.W. Bourque. 2001. The function of Ca2+ channel subtypes in exocytotic secretion: new perspectives from synaptic and non-synaptic release. Progress in Biophysics and Molecular Biology 77:269–303. Florey, E. 1954. An inhibitory and an excitatory factor of mammalian central nervous system, and their action of a single sensory neuron. Archives of Physiology and Biochemistry 62:33–53. Florey, E. 1991. GABA: history and perspectives. Canadian Journal of Physiology and Pharmacology 69:1049–1056. Frank, E. 1973. Matching of facilitation at the neuromuscular junction of the lobster: a possible case for influence of muscle on nerve. Journal of Physiology (London) 233:635–658. Franzini-Armstrong, C. 1976. Freeze-fracture of excitatory and inhibitory synapses in crayfish neuromuscular junctions. Journal de Microscopie et de Biologie Cellulaire 25:217–222. French, L.B., C.C. Lanning, and R.M. Harris-Warrick. 2002. The localization of two voltage-gated calcium channels in the pyloric network of the lobster stomatogastric ganglion. Neuroscience 112:217–232. Furshpan, E.J., and D.D. Potter. 1957. Mechanism of nerve-impulse transmission at a crayfish synapse. Nature 180:342–343. Furshpan, E.J., and D.D. Potter. 1959a. Transmission at the giant motor synapses of the crayfish. Journal of Physiology (London) 145:289–325.

169

170

Gregory Lnenicka Furshpan, E.J., and D.D. Potter. 1959b. Slow post-synaptic potentials recorded from the giant motor fibre of the crayfish. Journal of Physiology (London) 145:326–335. Govind, C.K., and R.G. Chiang. 1979. Correlation between presynaptic dense bodies and transmitter output at lobster neuromuscular terminals by serial section electron microscopy. Brain Research 161:377–388. Govind, C.K., and D.E. Meiss. 1979. Quantitative comparison of low- and high-output neuromuscular synapses from a motoneuron of the lobster (Homarus americanus). Cell and Tissue Research 198:455–463. Govind, C.K., J. Pearce, J.M. Wojtowicz, and H.L. Atwood. 1994. “Strong” and “weak” synaptic differentiation in the crayfish opener muscle: structural correlates. Synapse 16:45–58. Hashemzadeh-Gargari, H., and J.E. Freschi. 1992. Histamine activates chloride conductance in motor neurons of the lobster cardiac ganglion. Journal of Neurophysiology 68:9–15. Hatt, H., C. Franke, and J. Dudel. 1988. Calcium dependent gating of the L-glutamate activated, excitatory synaptic channel on crayfish muscle. Pflügers Archiv: European Journal of Physiology 411:17–26. He, P., R.C. Southard, D. Chen, S.W. Whiteheart, and R.L. Cooper. 1999. Role of α-SNAP in promoting efficient neurotransmission at the crayfish neuromuscular junction. Journal of Neurophysiology 82:3406–3416. He, T., and G.A. Lnenicka. 2011. Ca2+ buffering at a Drosophila larval synaptic terminal. Synapse 65:687–693. Heidelberger, R., C. Heinemann, E. Neher, and G. Matthews. 1994. Calcium dependence of the rate of exocytosis in a synaptic terminal. Nature 371:513–515. Heitler, W.J., A.H. Watson, S.W. Falconer, and B. Powell. 2001. Glutamate is a transmitter that mediates inhibition at the rectifying electrical motor giant synapse in the crayfish. Journal of Comparative Neurology 430:12–26. Hochner, B., H. Parnas, and I. Parnas. 1991. Effects of intra-axonal injection of Ca2+ buffers on evoked release and on facilitation in the crayfish neuromuscular junction. Neuroscience Letters 125:215–218. Hong, S.J., and G.A. Lnenicka. 1993. Long-term changes in the neuromuscular synapses of a crayfish motoneuron produced by calcium influx. Brain Research 605:121–127. Hong, S.J., and G.A. Lnenicka. 1995. Activity-dependent reduction in voltage-dependent calcium current in a crayfish motoneuron. Journal of Neuroscience 15:3539–3547. Hong, S.J., and G.A. Lnenicka. 1997. Characterization of a P-type calcium current in a crayfish motoneuron and its selective modulation by impulse activity. Journal of Neurophysiology 77:76–85. Hoyle, G., and C.A.G. Wiersma. 1958. Excitation at neuromuscular junctions in Crustacea. Journal of Physiology (London) 143:403–425. Hua, S.Y., M.A. Teylan, and A. Cimenser. 2007. An antibody to synaptotagmin I facilitates synaptic transmission. European Journal of Neuroscience 25:3217–3225. Jahromi, S.S., and H.L. Atwood. 1974. Three-dimensional ultrastructure of the crayfish neuromuscular apparatus. Journal of Cell Biology 63:599–613. Jeromin, A., A.J. Shayan, M. Msghina, J. Roder, and H.L. Atwood. 1999. Crustacean frequenins: molecular cloning and differential localization at neuromuscular junctions. Journal of Neurobiology 41:165–175. Johnson, B.R., J.H. Peck, and R.M. Harris-Warrick. 1994. Differential modulation of chemical and electrical components of mixed synapses in the lobster stomatogastric ganglion. Journal of Comparative Physiology A 175:233–249. Kamiya, H., and R.S. Zucker. 1994. Residual Ca2+ and short-term synaptic plasticity. Nature 371:603–606. Katz, P.S., M.D. Kirk, and C.K. Govind. 1993. Facilitation and depression at different branches of the same motor axon: evidence for presynaptic differences in release. Journal of Neuroscience 13:3075–3089. Kawagoe, R., K. Onodera, and A. Takeuchi. 1981. Release of glutamate from the crayfish neuromuscular junction. Journal of Physiology (London) 312:225–236. Kawai, N., and A. Niwa. 1977. Hyperpolarization of the excitatory nerve terminals by inhibitory nerve stimulation in lobster. Brain Research 137:365–368.



Synapses in Crustaceans

Kennedy, D., R.L. Calabrese, and J.J. Wine. 1974. Presynaptic inhibition: primary afferent depolarization in crayfish neurons. Science 186:451–454. King, M.J., H.L. Atwood, and C.K. Govind. 1996. Structural features of crayfish phasic and tonic neuromuscular terminals. Journal of Comparative Neurology 372:618–626. Krause, K.M., J. Pearce, and C.K. Govind. 1998. Regeneration of phasic motor axons on a crayfish tonic muscle: neuron specifies synapses. Journal of Neurophysiology 80:994–997. Kravitz, E.A., S.W. Kuffler, and D.D. Potter. 1963a. Gamma-aminobutyric acid and other blocking compounds in Crustacea. III. Their relative concentrations in separated motor and inhibitory axons. Journal of Neurophysiology 26:739–51. Kravitz, E.A., S.W. Kuffler, D.D. Potter, and N.M. Vangelder. 1963b. Gamma-aminobutyric acid and other blocking compounds in Crustacea. II. Peripheral nervous system. Journal of Neurophysiology 26:729–738. Kuffler, S.W., and C. Edwards. 1958. Mechanism of gamma aminobutyric acid (GABA) action and its relation to synaptic inhibition. Journal of Neurophysiology 21:589–610. Kuffler, S.W., and B. Katz. 1946. Inhibition at the nerve muscle junction in Crustacea. Journal of Neurophysiology 9:337–346. Le Bon-Jego, M., I. Masante-Roca, and D. Cattaert. 2006. State-dependent regulation of sensory-motor transmission: role of muscarinic receptors in sensory-motor integration in the crayfish walking system. European Journal of Neuroscience 23:1283–1300. Levi, R., and A.I. Selverston. 2006. Mechanisms underlying type I mGluR-induced activation of lobster gastric mill neurons. Journal of Neurophysiology 96:3378–3388. Lin, J.W., Q. Fu, and T. Allana. 2005. Probing the endogenous Ca2+ buffers at the presynaptic terminals of the crayfish neuromuscular junction. Journal of Neurophysiology 94:377–386. Lingle, C. 1980. The sensitivity of decapod foregut muscles to acetylcholine and glutamate. Journal of Comparative Physiology A 138:187–199. Lnenicka, G.A., and H.L. Atwood. 1985a. Age-dependent long-term adaptation of crayfish phasic motor axon synapses to altered activity. Journal of Neuroscience 5:459–467. Lnenicka, G.A., and H.L. Atwood. 1985b. Long-term facilitation and long-term adaptation at synapses of a crayfish phasic motoneuron. Journal of Neurobiology 16:97–110. Lnenicka, G.A., and H.L. Atwood. 1989. Impulse activity of a crayfish motoneuron regulated its neuromuscular synaptic properties. Journal of Neurophysiology 61:91–96. Lnenicka, G.A., and DeF. Mellon, Jr. 1983a. Changes in electrical properties and quantal current during growth of identified muscle fibres in the crayfish. Journal of Physiology (London) 345:261–284. Lnenicka, G.A., and DeF. Mellon, Jr. 1983b. Transmitter release during normal and altered growth of identified muscle fibres in the crayfish. Journal of Physiology (London) 345:285–296. Lnenicka, G.A., and E.J. Morley. 2002. Activity-dependent development and plasticity of crustacean motor terminals. Pages 266–281 in K. Wiese, editor. The crustacean nervous system. Springer-Verlag, Berlin Heidelberg. Lnenicka, G.A., H.L. Atwood, and L. Marin. 1986. Morphological transformation of synaptic terminals of a phasic motoneuron by long-term tonic stimulation. Journal of Neuroscience 6:2252–2258. Lnenicka, G.A., J.A. Blundon, and C.K. Govind. 1988. Early experience influences the development of bilateral asymmetry in a lobster motoneuron. Developmental Biology 129:84–90. Lnenicka, G.A., S.J. Hong, M. Combatti, and S. LePage. 1991. Activity-dependent development of synaptic varicosities at crayfish motor terminals. Journal of Neuroscience 11:1040–1048. Lnenicka, G.A., K.F. Arcaro, and J.M. Calabro. 1998. Activity-dependent development of calcium regulation in growing motor axons. Journal of Neuroscience 18:4966–4972. Magleby, K.L. 1973. The effect of tetanic and post-tetanic potentiation on facilitation of transmitter release at the frog neuromuscular junction. Journal of Physiology (London) 234:353–371. Magleby, K.L., and J.E. Zengel. 1976. Stimulation-induced factors which affect augmentation and potentiation of trasmitter release at the neuromuscular junction. Journal of Physiology (London) 260:687–717. Mallart, A., and A.R. Martin. 1967. An analysis of facilitation of transmitter release at the neuromuscular junction of the frog. Journal of Physiology (London) 193:679–694.

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Gregory Lnenicka Mamiya, A., and F. Nadim. 2005. Target-specific short-term dynamics are important for the function of synapses in an oscillatory neural network. Journal of Neurophysiology 94:2590–2602. Marder, E., and J.S. Eisen. 1984. Transmitter identification of pyloric neurons: electrically coupled neurons use different transmitters. Journal of Neurophysiology 51:1345–1361. Marder, E., and D. Paupardin-Tritsch. 1978. The pharmacological properties of some crustacean neuronal acetylcholine, gamma-aminobutyric acid, and L-glutamate responses. Journal of Physiology (London) 280:213–236. Marder, E., and D. Paupardin-Tritsch. 1980. The pharmacological profile of the acetylcholine response of a crustacean muscle. Journal of Experimental Biology 88:147–159. Marmont, G., and C.A.G. Wiersma. 1938. On the mechanism of inhibition and excitation of crayfish muscle. Journal of Physiology (London) 93:173–193. Matveev, V., A. Sherman, and R.S. Zucker. 2002. New and corrected simulations of synaptic facilitation. Biophysical Journal 83:1368–1373. Matveev, V., R.S. Zucker, and A. Sherman. 2004. Facilitation through buffer saturation: constraints on endogenous buffering properties. Biophysical Journal 86:2691–2709. McClintock, T.S., and B.W. Ache. 1989. Histamine directly gates a chloride channel in lobster olfactory receptor neurons. Proceedings of the National Academy of Sciences of the United States of America 86:8137–8141. Mercier, A.J., H. Bradacs, and H.L. Atwood. 1992. Long-term adaptation of crayfish neurons depends on the frequency and number of impulses. Brain Research 598:221–224. Millar, A.G., R.S. Zucker, G.C. Ellis-Davies, M.P. Charlton, and H.L. Atwood. 2005. Calcium sensitivity of neurotransmitter release differs at phasic and tonic synapses. Journal of Neuroscience 25:3113–3125. Miller, M.W., E.T. Vu, and F.B. Krasne. 1992. Cholinergic transmission at the first synapse of the circuit mediating the crayfish lateral giant escape reaction. Journal of Neurophysiology 68:2174–2184. Miwa, A., M. Ui, and N. Kawai. 1990. G protein is coupled to presynaptic glutamate and GABA receptors in lobster neuromuscular synapse. Journal of Neurophysiology 63:173–180. Miwa, A., H.P. Robinson, and N. Kawai. 1993. Presynaptic glutamate receptors depress inhibitory postsynaptic transmission in lobster neuromuscular synapse. Journal of Neurophysiology 70:1159–1167. Msghina, M., A.G. Millar, M.P. Charlton, C.K. Govind, and H.L. Atwood. 1999. Calcium entry related to active zones and differences in transmitter release at phasic and tonic synapses. Journal of Neuroscience 19:8419–8434. Mulkey, R.M., and R.S. Zucker. 1992. Posttetanic potentiation at the crayfish neuromuscular junction is dependent on both intracellular calcium and sodium ion accumulation. Journal of Neuroscience 12:4327–4336. Nagayama, T. 2005. GABAergic and glutamatergic inhibition of nonspiking local interneurons in the terminal abdominal ganglion of the crayfish. Journal of Experimental Zoology 303 A:66–75. Neher, E. 1995. The use of fura-2 for estimating Ca buffers and Ca fluxes. Neuropharmacology 34:1423–1442. Nguyen, P.V., and H.L. Atwood. 1990. Expression of long-term adaptation of synaptic transmission requires a critical period of protein synthesis. Journal of Neuroscience 10:1099–1109. Nguyen, P.V., and H.L. Atwood. 1997. Synaptic physiology and mitochondrial function in crayfish tonic and phasic motor neurons. Journal of Neurophysiology 78:281–294. Nguyen, P.V., L. Marin, and H.L. Atwood. 1994. Altered impulse activity modifies synaptic physiology and mitochondria in crayfish phasic motor neurons. Journal of Neurophysiology 72:2944–2955. Nusbaum, M.P., and M.P. Beenhakker. 2002. A small-systems approach to motor pattern generation. Nature 417:343–350. Ohta, Y., K. Nishikawa, Y. Hiroaki, and Y. Fujiyoshi. 2011. Electron tomographic analysis of gap junctions in lateral giant fibers of crayfish. Journal of Structural Biology 175:49–61. Onodera, K., and A. Takeuchi. 1975. Ionic mechanism of the excitatory synaptic membrane of the crayfish neuromuscular junction. Journal of Physiology (London) 252:295–318.



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Pahapill, P.A., G.A. Lnenicka, and H.L. Atwood. 1985. Asymmetry of motor impulses and neuromuscular synapses produced in crayfish claws by unilateral immobilization. Journal of Comparative Physiology A 157:461–467. Pahapill, P.A., G.A. Lnenicka, and H.L. Atwood. 1986. Neuronal experience modifies synaptic long-term facilitation. Canadian Journal of Physiology and Pharmacology 64:1052–1054. Pahapill P.A., G.A. Lnenicka and H.L. Atwood. 1987. Long-term facilitation and low-frequency depression in a crayfish phasic motor axon. Journal of Comparative Physiology A 161:367–375. Pan, B., and R.S. Zucker. 2009. A general model of synaptic transmission and short-term plasticity. Neuron 62:539–554. Pearce, J., C.K. Govind, and D.E. Meiss. 1985. Growth-related features of lobster neuromuscular terminals. Brain Research 353:215–228. Pearce, J., C.K. Govind, and R.R. Shivers. 1986. Intramembranous organization of lobster excitatory neuromuscular synapses. Journal of Neurocytology 15:241–252. Pearlstein, E., A.R. Marchand, and F. Clarac. 1994. Inhibitory effects of L-glutamate on central processes of crustacean leg motoneurons. European Journal of Neuroscience 6:1445–1452. Phelan, P. 2005. Innexins: members of an evolutionarily conserved family of gap-junction proteins. Biochimica et Biophysica Acta 1711:225–245. Pulver, S.R., D. Bucher, D.J. Simon, and E. Marder. 2005. Constant amplitude of postsynaptic responses for single presynaptic action potentials but not bursting input during growth of an identified neuromuscular junction in the lobster, Homarus americanus. Journal of Neurobiology 62:47–61. Rathmayer, W., S. Djokaj, A. Gaydukov, and S. Kreissl. 2002. The neuromuscular junctions of the slow and the fast excitatory axon in the closer of the crab Eriphia spinifrons are endowed with different Ca2+ channel types and allow neuron-specific modulation of transmitter release by two neuropeptides. Journal of Neuroscience 22:708–717. Robbins, J. 1959. The excitation and inhibition of crustacean muscle by amino acids. Journal of Physiology (London) 148:39–50. Rumpal, N., and G.A. Lnenicka. 2003. Ca2+ clearance at growth cones produced by crayfish motor axons in an explant culture. Journal of Neurophysiology 89:3225–3234. Ryan, R.M., and R.J. Vandenberg. 2005. A channel in a transporter. Clinical and Experimental Pharmacology and Physiology 32:1–6. Scanziani, M., B.H. Gahwiler, and S. Charpak. 1998. Target cell-specific modulation of transmitter release at terminals from a single axon. Proceedings of the National Academy of Sciences of the United States of America 95:12004–12009. Schneggenburger, R., and E. Neher. 2005. Presynaptic calcium and control of vesicle fusion. Current Opinion in Neurobiology 15:266–274. Sherff, C.M., and B. Mulloney. 1996. Tests of the motor neuron model of the local pattern-generating circuits in the swimmeret system. Journal of Neuroscience 16:2839–2859. Sherman, R.G., and H.L. Atwood. 1971. Synaptic facilitation: long-term neuromuscular facilitation in crustaceans. Science 171:1248–1250. Sherman, R.G., and H.L. Atwood. 1972. Correlated electrophysiological and ultrastructural studies of a crustacean motor unit. Journal of General Physiology 59:586–615. Shinozaki, H. 1980. The pharmacology of the excitatory neuromuscular junction in the crayfish. Progress in Neurobiology 14:121–155. Shupliakov, O., H.L. Atwood, O.P. Ottersen, J. Storm-Mathisen, and L. Brodin. 1995. Presynaptic glutamate levels in tonic and phasic motor axons correlate with properties of synaptic release. Journal of Neuroscience 15:7168–7180. Silverman-Gavrila, L.B., and M.P. Charlton. 2009. Calcineurin and cytoskeleton in low-frequency depression. Journal of Neurochemistry 109:716–732. Silverman-Gavrila, L.B., P.M. Orth, and M.P. Charlton. 2005. Phosphorylation-dependent low-frequency depression at phasic synapses of a crayfish motoneuron. Journal of Neuroscience 25:3168–3180. Takeuchi, A., and N. Takeuchi. 1964. The effect on crayfish muscle of iontophoretically applied glutamate. Journal of Physiology (London) 170:296–317.

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Gregory Lnenicka Takeuchi, A., and N. Takeuchi. 1965. Localized action of gamma-aminobutyric acid on the crayfish muscle. Journal of Physiology (London) 177:225–238. Takeuchi, A., and N. Takeuchi. 1966. On the permeability of the presynaptic terminal of the crayfish neuromuscular junction during synaptic inhibition and the action of gamma-aminobutyric acid. Journal of Physiology (London) 183:433–449. Tank, D.W., W.G. Regehr, and K.R. Delaney. 1995. A quantitative analysis of presynaptic calcium dynamics that contribute to short-term enhancement. Journal of Neuroscience 15:7940–7952. Tisdale, A.D., and Y. Nakajima. 1976. Fine structure of synaptic vesicles in two types of nerve terminals in crayfish stretch receptor organs: influence of fixation methods. Journal of Comparative Neurology 165:369–386. Turrigiano, G.G., and S.B. Nelson. 2004. Homeostatic plasticity in the developing nervous system. Nature Reviews Neuroscience 5:97–107. Uchizono, K. 1967. Inhibitory synapses on the stretch receptor neurone of the crayfish. Nature 214:833–834. van Harreveld, A., and M. Mendelson. 1959. Glutamate-induced contractions in crustacean muscle. Journal of Cellular and Comparative Physiology 54:85–94. Velez, S.J., and R.J. Wyman. 1978a. Synaptic connectivity in a crayfish neuromuscular system. I. Gradient of innervation and synaptic strength. Journal of Neurophysiology 41:75–84. Velez S.J., and R.J. Wyman. 1978b. Synaptic connectivity in a crayfish neuromuscular system. II. Nerve-muscle matching and nerve branching patterns. Journal of Neurophysiology 41:85–96. Watanabe, A. 1958. The interaction of electrical activity among neurons of lobster cardiac ganglion. Japanese Journal of Physiology 8:305–318. Wojtowicz, J.M., and H.L. Atwood. 1985. Correlation of presynaptic and postsynaptic events during establishment of long-term facilitation at crayfish neuromuscular junction. Journal of Neurophysiology 54:220–230. Wojtowicz, J.M., and H.L. Atwood. 1986. Long-term facilitation alters transmitter releasing properties at the crayfish neuromuscular junction. Journal of Neurophysiology 55:484–498. Wojtowicz, J.M., and H.L. Atwood. 1988. Presynaptic long-term facilitation at the crayfish neuromuscular junction: voltage-dependent and ion-dependent phases. Journal of Neuroscience 8:4667–4674. Wojtowicz, J.M., L. Marin, and H.L. Atwood. 1994. Activity-induced changes in synaptic release sites at the crayfish neuromuscular junction. Journal of Neuroscience 14:3688–3703. Yamada, W.M., and R.S. Zucker. 1992. Time course of transmitter release calculated from simulations of a calcium diffusion model. Biophysical Journal 61:671–682. Zhong, N., V. Beaumont, and R.S. Zucker. 2001. Roles for mitochondrial and reverse mode Na+/Ca2+ exchange and the plasmalemma Ca2+ ATPase in post-tetanic potentiation at crayfish neuromuscular junctions. Journal of Neuroscience 21:9598–9607. Zucker, R.S. 1972. Crayfish escape behavior and central synapses. I. Neural circuit exciting lateral giant fiber. Journal of Neurophysiology 35:599–620. Zucker, R.S. 1974. Characteristics of crayfish neuromuscular facilitation and their calcium dependence. Journal of Physiology (London) 241:91–110. Zucker, R.S., and J. Bruner. 1977. Long-lasting depression and the depletion hypothesis at crayfish neuromuscular junctions. Journal of Comparative Physiology A 121:223–240. Zucker, R.S., and W.G. Regehr. 2002. Short-term synaptic plasticity. Annual Review of Physiology 64:355–405.

7 ADULT NEUROGENESIS IN CRUSTACEANS

Manfred Schmidt

Abstract In decapod crustaceans, adult neurogenesis—the generation of new neurons in an adult animal— occurs in sense organs (compound eyes and sensilla) and the brain. Generation of new receptor neurons of sense organs is linked to molting underlying the indeterminate growth of adults and it occurs as addition of entire sensory units (ommatidia or sensilla) to preexisting arrays. Adult neurogenesis of receptor neurons has been analyzed for aesthetascs (olfactory sensilla). New olfactory receptor neurons (ORNs) are generated in a proliferation zone proximal to the array of functionally mature aesthetascs, and old ORNs are lost distally with molting. This generates a slow turnover of ORNs and results in a longitudinal age gradient along the aesthetasc array. In the brain, new interneurons are generated in the optic lobes and the central olfactory pathway. While adult neurogenesis in the optic lobes appears to be linked to the addition of ommatidia to the compound eyes, adult neurogenesis in the central olfactory pathway is a continuous process independent of the addition of new ORNs. Adult neurogenesis in the central olfactory pathway occurs focally in small proliferation zones located deep within the clusters of somata of mature interneurons. All decapod species analyzed have adult neurogenesis in the olfactory deutocerebrum, which includes the first synaptic relay of the central olfactory pathway, the olfactory lobe. In the olfactory deutocerebrum, each proliferation zone is associated with one large adult neuroblast, likely acting as an asymmetrically dividing neural stem cell. Each adult neuroblast is embedded in a presumptive neurogenic niche composed of small, bipolar cells, whose identity is currently controversial (glial vs. ectodermal). Overall adult neurogenesis in the olfactory deutocerebrum of decapods parallels adult neurogenesis in the brain of mammals and insects in being based on the same type of neuronal stem cell as is embryonic neurogenesis.

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INTRODUCTION The developmental process leading to the generation of new neurons in the nervous system is called neurogenesis. In the strictest sense, this comprises the birth of an immature neuron through the cell division of a precursor cell, its differentiation, and morphological and physiological maturation. More often the term “neurogenesis” is used in a broader sense including the processes that lead to the generation of the neural precursor cells, and this broader definition will be used throughout this review. Common to all Bilateria is that their nervous system is of ectodermal origin and that it consists of a central nervous system (CNS) and a peripheral nervous system (PNS). Across Bilateria, the first wave of neurogenesis occurs during embryogenesis, and it begins with the specification of a neuroectoderm or neuroepithelium from previously undifferentiated ectodermal cells (Egger et al. 2008, Stollewerk 2008, Kriegstein and Alvarez-Buylla 2009, Meyer and Seaver 2009). In vertebrates and Tetraconata (the likely monophyletic taxon comprising Hexapoda and Crustacea: Dohle 2001, Regier et al. 2010), the embryonic CNS is generated by neural stem cells (NSCs) arising from the neuroectoderm. Neural stem cells fuel embryonic neurogenesis through a series of cell divisions, at least some of which are asymmetric, generating one daughter that is the self-renewed NSC and another daughter that is destined to become or to produce differentiated neurons. In other bilaterian taxa such as myriapods, chelicerates, and nematodes, embryonic neurogenesis proceeds without the involvement of NSCs (Stollewerk 2008). In most Bilateria, neurogenesis does not cease after embryogenesis but continues further, often throughout life. Thus neurogenesis typically is a prolonged process spanning diverse developmental stages (embryo, larva, juvenile, and adult) with distinct demands reflected by specific stem cell properties (He et al. 2009). Clearly distinguishing these developmental stages is a basic requirement for analyzing neurogenesis in each of them (Rakic 2002). In particular, this applies to adult neurogenesis, which is defined as neurogenesis in the PNS or CNS of an adult animal, as it hinges on establishing a criterion for adulthood. The commonly agreed on biological indicator for adulthood is sexual maturity (Lindsey and Tropepe 2006), and in animals in which reaching sexual maturity is accompanied by distinct morphological changes (e.g., imaginal molt of insects, pubertal molt of some decapod crustaceans), adulthood can readily be established. However, in many other taxa, including most vertebrates and decapod crustaceans, the transition from the juvenile to the adult stage is morphologically indistinct and thus the experimental animals’ age or size is regularly used as determinant of adulthood (Hartnoll 1985). Using these criteria for adulthood, adult neurogenesis frequently occurs in the PNS and CNS of crustaceans and vertebrates and in the CNS of insects (Lindsey and Tropepe 2006). Adult neurogenesis may occur in other invertebrate taxa (e.g., Packard and Albergoni 1970), but these have not been assayed systematically. Since the mid-1990s, the study of adult neurogenesis has gained enormous popularity mainly for two reasons: (1) At that time, adult neurogenesis was established as a constitutive process in the brain of mammals including humans (Gross 2000), raising the hope that understanding its cellular and molecular underpinnings could aid in the discovery of treatments for degenerative brain disorders or traumatic brain injuries (Curtis et al. 2003). (2) Methodological progress led to the development of immunocytochemical techniques for labeling proliferating cells in vivo with 5-bromo-2’-deoxyuridine (BrdU) or other thymidine analogs that are incorporated into newly synthesized DNA during the S-phase of the cell cycle (Fig. 7.1A) (Taupin 2007). In vivo labeling with BrdU enabled proliferating cells to be readily located within nervous tissue and studied with respect to their cellular makeup and fate. However, demonstration of BrdU+ cells in neuronal tissue is insufficient to demonstrate neurogenesis, since BrdU can also be incorporated into DNA during DNA repair or endoreplication, as a prelude to apoptosis, and proliferating cells can give rise to nonneuronal cell types such as glial cells (Rakic 2002, Taupin 2007).



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Fig. 7.1. Cell cycle and S-phase labeling with BrdU. (A) The cell cycle has four phases (G1, S, G2, M) the duration of which is represented by the length of the corresponding arrow. BrdU labels cells in S-phase by incorporation into newly synthesized DNA, and anti-pH3 labels cells in M-phase at the time of fixation. (B) Schematic depiction of influence of S-phase duration on the number of cells labeled by BrdU. Symbolized are two populations of 10 cycling cells that differ in S-phase duration (gray bars) but are identical in the proliferation rate (all cells divide once in the depicted time interval). Dark gray bar on top represents the time BrdU is available for incorporation into DNA. Note that 8 of 10 cells with a long S-phase but only 2 of 10 cells with a short S-phase are labeled by BrdU (dark gray); unlabeled cells (light gray).

Therefore, BrdU labeling must be combined with other methods to demonstrate that cells are newly generated and then differentiate into neurons. In the mid- to late 1990s, adult neurogenesis was for the first time demonstrated in three invertebrate neuronal systems: (1) the brain of insects, where in some species new Kenyon cells, the intrinsic neurons constituting the mushroom bodies, are produced in adults (see review by Cayre et al. 2007); (2) the ORNs of decapod crustaceans (Sandeman and Sandeman 1996, Steullet et al. 2000); and (3) the brain of decapod crustaceans, where new neurons are continuously generated in the central olfactory pathway (Schmidt 1997, Sandeman et al. 1998, Schmidt and Harzsch 1999).

CRUSTACEAN LIFE HISTORY AND EMBRYONIC/LARVAL NEUROGENESIS To understand adult neurogenesis in the PNS and CNS of decapod crustaceans in its organismal context, it is necessary to give a short overview of the life history of these animals and of what is known about embryonic and larval neurogenesis in Tetraconata. The terms “decapod crustaceans” or “decapods” will be used for the crustacean order Decapoda. For taxa within Decapoda, the following terms (based on Martin and Davis 2001) will be used: “shrimp” (superfamily Penaeoidea and infraorders Caridea and Stenopodidea), “Reptantia” (all decapods other than shrimp), “spiny lobsters” (family Palinuridae), “slipper lobsters” (family Scyllaridae), “clawed lobsters” (superfamily Nephropidae), “crayfish” (superfamily Astacoidea), “anomuran crabs” (infraorder Anomura), “brachyuran crabs” (infraorder Brachyura). Life History of Decapods—Indeterminate Adult Growth Crustaceans are covered by a hardened exoskeleton (cuticle) and grow discontinuously through a series of molts in which the old cuticle is shed. This generates a series of intermolt phases

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Fig. 7.2. Lifelong growth of decapods and principles of neurogenesis in PNS and CNS. (A) Artist’s rendering of an early juvenile and an old adult Carcinus maenas, highlighting the dramatic lifelong growth (from Schmidt 1997, with permission from Elsevier Limited). (B) Addition of sensilla to the telson of Procambarus clarkii from POI hatchlings (top) to adults (bottom); scale bar = 200 µm (from Letourneau 1976, with permission from Springer). (C)  Construction of sensilla in crustaceans (top) and insects (bottom) (from Schmidt and Gnatzy 1984, with permission from Springer). Mechanoreceptor neurons (dark gray), chemoreceptor neurons (light gray). Abbreviations: cS, ciliary segment of outer dendrite; iD, inner dendritic segment; R, rootlet; Sc, scolopale; spC, spongy cuticle; tP, terminal pore; WP, wall pores. (D)  Model of embryonic neurogenesis in crustaceans. 1, 2. Bipolar neuroectodermal cells (light gray) likely connected apically by adherens junctions (dark gray boxes) multiply by symmetrical divisions in the ectodermal plane (white arrows), and some of them differentiate into large, globose neuroblasts (NB; black arrow). 3, 4. NBs undergo self-renewing asymmetric divisions oriented perpendicularly to the ectoderm and bud off smaller ganglion mother cells (GMC) into the body. 4, 5. GMCs divide once symmetrically and produce two immature neurons (iN).

or instars, and if there is a significant change in morphology between two instars this molt is called a metamorphosis (Williamson 1982). Most decapods have indirect development; they hatch as a larva with a morphology that is substantially different from the adult, and subsequent development involves one or more metamorphoses. Larval development is terminated by the metamorphosis to the first juvenile with an adult-like morphology. Some decapods, including all crayfishes, have a direct development; they begin their postembryonic free life as juvenile with an adult-like morphology (Williamson 1982). Thus, the life history of decapods comprises three or four distinct developmental stages: (1) embryogenesis terminated by hatching, (2) larval development (in animals with indirect development) often with several morphologically distinct stages, (3) juvenile development with gradual differentiation of sexual characters, and



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(4) adult life after reaching sexual maturity. In most decapods, the transition from juvenile to sexually mature adult is not accompanied by overt changes in outer morphology, but in some brachyuran and anomuran crabs as well as in many shrimp, this transition is through a “pubertal molt” involving distinct morphological changes (Hartnoll 1985). Most decapods continue to grow indeterminately as adults, albeit with a decreasing frequency of molting and decreasing growth increment per molt (Hartnoll 1985). Some brachyuran crabs have a “terminal molt” after which no further molting or growth occurs, and in some species (especially spider crabs) the pubertal molt of females is their terminal molt (Carlisle 1957, Hartnoll 1985). As adults, decapods have a typical life expectancy in the range of several years to some decades (Farmer 1973, Sheehy et al. 1999, Maxwell et al. 2007). From the first juvenile stage until the animals reach their maximal adult size, decapods grow enormously (Fig. 7.2A). The length or width of the body typically increases between ~10- and 100-fold, which amounts to a ~100to 10,000-fold increase in surface area (Schmitz 1992) and a ~1000- to 1,000,000-fold increase in body volume (= weight). The clawed lobster Homarus americanus, for instance, increases ~300,000-fold in weight from 60 mg in fifth instar juveniles to > 19 kg in the largest adults on record (Wolff 1978, Lavalli 1990). Embryonic and Larval Neurogenesis in Tetraconata To understand neurogenesis in the PNS and CNS of adult decapods, it is instructive to know how neurons are generated during embryonic and larval development of crustaceans and insects: (1) In crustaceans and insects, embryonic and larval neurogenesis in the CNS is maintained by likely homolog NSCs called neuroblasts (NBs) (Duman-Scheel and Patel 1999, Harzsch 2003, Ungerer and Scholtz 2008, Ungerer et al. 2011). (2) In crustaceans and insects, the neuronal systems of the CNS and the PNS that are formed through neurogenesis are highly similar in morphological layout (Fig. 7.2C) (Schmidt and Gnatzy 1984, Dohle 2001, Schachtner et al. 2005, Strausfeld 2009). (3) In insects, the morphological processes and molecular mechanisms involved in neurogenesis are known in great detail, and this can inform about equivalent processes in the less well-studied crustaceans (Keil 1997, Hartenstein 2005, Egger et al. 2008, Hartenstein et al. 2008). Embryonic and Larval Neurogenesis in the Central Nervous System of Tetraconata Among crustaceans, embryonic and larval neurogenesis in the CNS has been analyzed mainly in malacostracans, including decapods and a few branchiopods. In each case, neurogenesis is maintained by large, spherical NBs that differentiate from the ventral ectoderm (Fig. 7.2D) (Dohle 1976, Scholtz 1992, Gerberding 1997, Harzsch et al. 1999, Ungerer and Scholtz 2008). In decapods and most other malacostracans, NBs arise through invariant lineages from ectodermal stem cells (ectoteloblasts) and, in repetitive asymmetric divisions oriented perpendicularly to the body surface, they self-renew and bud off a smaller daughter cell, a ganglion mother cell (GMC), at their inward-facing pole. Typically, small cell clones consisting of a NB and some GMCs generated by it stay together and form an inward-facing cell column. Eventually, each GMC undergoes one terminal, symmetric division in which two postmitotic cells are produced that can be either immature neurons or glia. In the early phase of postembryonic development, neurogenesis in the CNS of decapods continues in the embryo-typical fashion with large NBs and smaller GMCs forming small, discrete clusters of proliferating cells (Harzsch and Dawirs 1994, 1995/96, 1996, Song et al. 2009). Embryonic and larval neurogenesis in the insect CNS is also maintained by large spherical NBs that through a series of asymmetric divisions self-renew and bud off smaller GMCs,

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Embryonic and Larval Neurogenesis in the Peripheral Nervous System of Tetraconata The main components of the PNS of Tetraconata are two types of sense organs, compound eyes and sensilla. Compound eyes are composed of many ommatidia (facets) each containing eight bipolar photoreceptor neurons and 11 accessory cells (Nilsson and Kelber 2007). Sensilla are small epidermal organules (Kleinorgane: Henke 1953) that occur on all body parts and are innervated by 1–500 bipolar receptor neurons (Keil and Steinbrecht 1984, Schmidt and Gnatzy 1984, Hallberg and Skog 2011, Schmidt and Mellon 2011, see also ­chapters 8 and 9 in this volume). Sensilla are subdivided into cuticular sensilla, which have an external cuticular apparatus (most often a hairor bristle-like seta), and scolopidia, which lack an external cuticular structure and typically form larger aggregates called chordotonal organs (Schmidt and Gnatzy 1984, Field 2005). In holometabolous insects, the compound eyes develop during larval stages from eye-antennal imaginal disks originating from small, invaginated pieces of embryonic blastoderm, and in hemimetabolous insects, they develop in the embryo from the ectoderm of optic primordia and continue to grow by addition of ommatidia through juvenile stages (Ready et  al. 1976, Morante et  al. 2007). The cells making up an ommatidium, including the photoreceptor neurons, are generated by one or two mitotic waves sweeping across the epithelium. Their determination is not by cell lineage, but through a process of sequential recruitment along a morphogenetic front (Ready et al. 1976, Morante et al. 2007). In crustaceans, the compound eyes develop in the embryo from the ectoderm of optic primordia by a proliferative and morphogenetic process that shows similarities to insects (Melzer et al. 2000, Hafner and Tokarski 2001). In many species, including decapods, the compound eyes continue to grow through larval and juvenile stages by addition of ommatidia to the outer margins of the eye (e.g., Parker 1890, Eguchi et al. 1989, Harzsch and Dawirs 1995/96, Melzer et al. 2000, Hafner and Tokarski 2001). Each sensillum of Tetraconata is composed of a specific complement of bipolar receptor neurons and several auxiliary cells (Fig. 7.2C) (Keil and Steinbrecht 1984, Schmidt and



Adult Neurogenesis in Crustaceans

Gnatzy 1984, Field 2005, Hallberg and Skog 2011, Schmidt and Mellon 2011, see c­ hapters 3 and 10 in this volume). The receptor neurons have an apical ciliated dendrite and a basal axon projecting to the CNS. Each receptor neuron serves a discrete sensory modality. Unimodal sensilla (e.g., olfactory sensilla, scolopidia) contain receptor neurons of the same modality, and bimodal sensilla (e.g., mechano- and chemoreceptive sensilla) contain specific complements of receptor neurons of different modalities. Across Tetraconata, some sensilla are generated in the embryo. At each subsequent molt during larval and juvenile development, new sensilla are added while the existing ones undergo a molting process in which their old cuticular apparatus (in cuticular sensilla) is replaced (e.g., Sandeman and Sandeman 1996, Steullet et al. 2000). Thus, neurogenesis of receptor neurons in the PNS of Tetraconata occurs during embryogenesis and continues throughout postembryonic development. In crustaceans, the generation of new sensilla has rarely been studied, and it is unknown how the receptor neurons and auxiliary cells arise (Guse 1983, Ekerholm and Hallberg 2002). In insects, the cell lineages and molecular signaling mechanisms underlying sensilla formation have been analyzed in detail (Keil 1997, Hartenstein 2005). Typically, each sensillum arises from one “sense organ precursor” (SOP), an ectodermal cell singled out from surrounding ectodermal cells in a selection process involving proneural genes from the family of helix-loop-helix (bHLH) proteins (Hartenstein 2005). The SOP divides in a non-stem-cell-like fashion and gives rise to the sensillum-specific complement of receptor neurons and auxiliary cells through a sensillum-specific, invariant cell lineage (Hartenstein 2005).

JUVENILE AND ADULT NEUROGENESIS IN DECAPODS Juvenile and Adult Neurogenesis in the Peripheral Nervous System Neurogenesis of receptor neurons in the PNS of juvenile and adult decapods is well documented for compound eyes and sensilla. Typically, the generation of new receptor neurons occurs in the context of growth-related addition of entire morphological units (ommatidia, sensilla) through molting. Only in specific sensilla (aesthetascs, see below) are new receptor neurons also added to young sensilla until the mature complement of receptor neurons is reached (Ekerholm and Hallberg 2002). Overall, the continuous addition of new receptor neurons throughout juvenile and adult life causes a continuous and substantial increase in the number of receptor neuron axons innervating the CNS. In H.  americanus, for instance, the number of axons of receptor neurons located in the chelae increases from ~20,000 in fifth instar juveniles to ~1,000,000 in large adults (Govind and Potter 1987). New ommatidia are continuously added throughout juvenile and adult life at the margin of the compound eye of diverse decapods (e.g., Bernhards 1916, Eguchi et al. 1989, Meyer-Rochow et  al. 1990), implying that juvenile and adult neurogenesis of photoreceptor neurons exists. However, it is unknown how this occurs. New sensilla of diverse types are added to existing arrays of sensilla on various appendages of decapods throughout juvenile and adult life. These include (1) diverse sensilla on the tail fan (Fig. 7.2B) (Letourneau 1976, Schmitz 1992, Stuart and Macmillan 1997); (2)  short mechanoreceptive sensilla forming “cuticular articulated peg” organs at joints of walking legs (Laverack 1976, 1978, Macmillan et  al. 1998); (3)  mechanoreceptive sensilla in the statocyst (Finley and Macmillan 2000); (4)  bimodal, mechano- and chemoreceptive sensilla (hedgehog hairs) on the walking leg chelae (Laverack 1988); (5) diverse types of sensilla (including aesthetascs, see below) located on the lateral flagellum of the first antennae, or antennules

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Manfred Schmidt (e.g., Kamiguchi 1972, Farmer 1973, Mellon et al. 1989, Sandeman and Sandeman 1996, Steullet et al. 2000, Harrison et al. 2001a, 2001b, Derby et al. 2003); and (6) scolopidia composing the propodite-dactylopodite (PD) chordotonal organ (Cooper and Govind 1991, Hartman and Cooper 1994). Juvenile and Adult Neurogenesis of Olfactory Receptor Neurons Only in aesthetascs, the olfactory sensilla of crustaceans located on the lateral flagella of the antennules (Hallberg and Skog 2011, Schmidt and Mellon 2011), have the developmental processes leading to the generation of new receptor neurons during juvenile and adult life been studied (Fig. 7.3A–D). Decapod aesthetascs are innervated by 40–500 olfactory receptor neurons (ORNs) whose somata typically form a discrete, subepithelial cluster below the associated tube-shaped seta, which has a very thin and permeable cuticle. The bundle of dendrites arising from the cluster of ORN somata is surrounded by numerous auxiliary cells (e.g., Grünert and Ache 1988). The lateral flagellum of the antennule on which the aesthetascs reside is annulated,

Fig. 7.3. Adult neurogenesis in the PNS of decapods. (A–C) Life-long addition of aesthetascs in Panulirus argus (from Harrison et al. 2001b, with permission from John Wiley and Sons). (A) Growth-related increase in the number of aesthetasc-bearing and other annuli of the lateral flagellum throughout juvenile and adult life; inset: SEM micrograph of tuft region of the lateral flagellum, aesthetascs (arrow); scale bar = 200 µm. (B) Proximal proliferation zone revealed by in vivo BrdU labeling. Numerous BrdU-positive cells are located within newly generated clusters of ORNs (1: single arrow; BrdU-positive cells outside clusters: double arrows) and at the bundles of inner dendrites where auxiliary cells reside (2: single arrows). (C) Schematic overview of ORN development. Black dots (BrdU+ cells). (D) Growth-related increase in the number of aesthetascs throughout juvenile and adult life in Cherax destructor (from Sandeman et al. 1998, with permission from The Society for Neuroscience). Inset: SEM micrograph of aesthetasc-bearing annulus of the lateral flagellum, aesthetascs (arrow); scale bar = 25 µm (from Sandeman and Sandeman 1996, with permission from The Company of Biologists).



Adult Neurogenesis in Crustaceans

but not all annuli bear aesthetascs. Typically, aesthetascs are restricted to a certain region where they constitute a tuft. Aesthetascs are accompanied by other sensilla (asymmetric, guard, and companion setae) with morphological features of bimodal, chemo- and mechanoreceptive sensilla (Hallberg and Skog 2011, Schmidt and Mellon 2011). In several species of decapods (including the crayfish Procambarus clarkii and Cherax destructor and the spiny lobster Panulirus argus), it has been shown that the increase in the number of aesthetascs throughout juvenile and adult life is due to the addition of aesthetasc-bearing annuli (Fig. 7.3A,D) (Kamiguchi 1972, Farmer 1973, Mellon et al. 1989, Sandeman and Sandeman 1996, Derby et al. 2003). Closer analysis of this process in C. destructor (Sandeman and Sandeman 1996) and P. argus (Steullet et al. 2000, Harrison et al. 2001a, 2001b, Derby et al. 2003) revealed that with each molt new aesthetasc-bearing annuli are added proximal to the tuft region. In adults, there is also molt-related loss of aesthetascs and annuli at the tip of the lateral flagellum. Since fewer annuli and aesthetascs are lost distally than are generated proximally, the number of annuli and aesthetascs increases continuously with animal size. Overall, this mode of growth creates a longitudinal age gradient along the tuft region with the youngest annuli and sensilla located proximally and the oldest ones distally. In adults, the additional distal loss of annuli, aesthetascs, and aesthetasc-associated sensilla creates a slow, longitudinal turnover of these sensory units. For young adult P. argus, the life span of a particular aesthetasc is in the range of 3–6 molt cycles amounting to ~6–12 months (Steullet et al. 2000). In vivo labeling with BrdU revealed that in adult P. argus new aesthetascs are formed in a continuous process within a restricted region, the “proximal proliferation zone” (PPZ) (Fig. 7.3B,C) (Harrison et al. 2001b). The PPZ is ~10 annuli long and includes some of the proximal-most aesthetasc-bearing annuli and several of the distal-most, non-aesthetasc-bearing annuli. The generation of aesthetascs occurs in a molt-stage-dependent spatiotemporal wave of proliferation and differentiation that travels proximolaterally from the array of preexisting aesthetascs through the PPZ. The generation of a new aesthetasc starts with the formation of a patch of proliferating epithelial cells expressing a spiny lobster achaete-scute homolog (Splash) (Chien et al. 2009, Tadesse et al. 2011). From the epithelial patch, a small cluster of subepidermal cells develops and expands through continuous proliferation of at least three types of precursor cells, generating primordial ORNs, auxiliary cells, and axon-associated glial cells. Finally, the seta is formed late in the premolt phase. After molting, the newly generated aesthetascs functionally mature indicated by a gradual acquisition of odor responsiveness within the ORN clusters (Steullet et al. 2000). Functional maturation takes several months and is not synchronized between the ORNs of an aesthetasc. The development of aesthetascs and insect olfactory sensilla shows some similarities. As in insect olfactory sensilla, the aesthetascs appear to originate from multiple precursors that may not be clonally related (Reddy et al. 1997), and all cells of the developing sensillum proliferate at the same time. Adult Neurogenesis in the Central Nervous System Organization of the Central Nervous System and of the Central Olfactory Pathway within the Brain The crustacean CNS consists of a series of segmental ganglia that are organized into central neuropils comprising dendritic and axonal processes of neurons and peripheral clusters of neuronal somata (Sandeman et al. 1992). Besides neurons, the CNS contains glial cells of several morphologically distinct types and perivascular cells forming the walls of arteries and arterioles, which constitute a highly branched vascular system within the CNS; and hemocytes (Abbott 1971, Cuadras and Marti-Subirana 1987, Schmidt and Derby 2011). In the anterior body region, the

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Fig. 7.4. Adult neurogenesis in the brain of Carcinus maenas. (A) Location of neurogenic regions in the central olfactory pathway (modified from Schmidt 2007b, with permission from Oxford University Press). Neuronal soma clusters containing proliferation zones (PZ) are highlighted in light gray. Abbreviations: A I Nv, antennular nerve; C, chiasm; HB, hemiellipsoid body; HBC, hemiellipsoid body cluster; LC, lateral soma cluster; MC, medial soma cluster; MT, terminal medulla; EC, esophageal connective; OGT, olfactory globular tract; OL, olfactory lobe; OPL, optic lobe; PT, protocerebral tract. (B and C) Proliferation zones revealed by in vivo BrdU labeling (modified from Schmidt 1997, with permission from Elsevier Limited). (B) Central brain. Groups of BrdU+ cells are located in PZs in the LCs (arrowheads), additional singular BrdU+ cells (arrows) occur in all brain compartments; inset: PZ of LC at higher magnification. (C) Eyestalk ganglion. The neuronal soma cluster (asterisk) at the hemiellipsoid body (HB) contains a proliferation zone with a dense group of BrdU+ cells; inset: PZ at higher magnification. (D) Growth-related increase in the number of projection neurons (somata, axons) in the olfactory deutocerebrum throughout juvenile and adult life (from Schmidt 1997, with permission from Elsevier Limited). (E) Number of BrdU+ cells in the proliferation zone of the LC at different survival times after BrdU injection. The number of BrdU+ cells stays constant up to a survival time of five days but approximately doubles at a survival time of one month (from Schmidt 1997, with permission from Elsevier Limited). (F) Number of BrdU+ cells in the proliferation zone of the LC in crabs of different sizes, either directly after their catch or after being held in captivity for 6 weeks. The number of BrdU+ cells decreases with animal size, and it is lower after crabs were held in captivity (from Hansen and Schmidt 2004, with permission from Elsevier Limited).



Adult Neurogenesis in Crustaceans

Fig. 7.5. Organization of proliferation zones in the olfactory deutocerebrum of adult Panulirus argus. (A) Location of neurogenic complexes in the central brain. Neuronal soma clusters containing proliferation zones (PZ) are highlighted in light gray (from Schmidt and Derby 2011, with permission from John Wiley and Sons). Abbreviations: A I Nv, antennular nerve; AL, accessory lobe; LC, lateral soma cluster; LN, local interneuron; MC, medial soma cluster; MP, median protocerebrum; NN, neurogenic niche; OGT, olfactory globular tract; OL, olfactory lobe; ORN, olfactory receptor neuron; PN, projection neuron; PT, protocerebral tract. (B) Proliferation zones in the olfactory deutocerebrum (one hemibrain) revealed by in vivo BrdU labeling; insets: respective PZs at higher magnification. Abbreviations: AL, accessory lobe; OL, olfactory lobe; scale bar = 500 µm (modified from Schmidt and Harzsch 1999, with permission from Marine Biological Laboratory). (C) Methylene blue-stained semithin sections through the LC shows PZ composed of small, darkly stained cells contrasting with neuronal somata (N) and containing a cell in mitosis (arrow); scale bar = 50 µm (modified from Schmidt 2001, with permission from John Wiley and Sons). (D) MC triple-labeled with the nuclear marker Hoechst (light gray) and antibodies against the neuropeptides FMRFamide and Substance P (white). The cells composing the PZ are FMRF- and SP-negative and hence nonneuronal; scale bar = 50 µm (modified from Schmidt 2001, with permission from John Wiley and Sons). (E) TEM micrograph of the center of the PZ in the LC. The PZ is composed of type A cells (PZ-A), type B cells (PZ-B), and type C cells (PZ-C); scale bar = 5 µm (from Schmidt and Derby 2011, with permission from John Wiley and Sons).

supraesophageal ganglion or brain is formed by three fused segmental neuromeres: the protocerebrum, receiving input from the compound eyes; the deutocerebrum, receiving input from the antennules; and the tritocerebrum, receiving input from the second antennae (Sandeman et al. 1992, see ­chapter 2 in this volume). In most decapods, the brain is structured into three large compartments:  the central brain located medially within the head and an eyestalk ganglion in each of both eyestalks connected with the central brain via a protocerebral tract (PT) (Fig. 7.4A). The eyestalk ganglion contains the optic lobe and the lateral protocerebrum composed of hemiellipsoid body and terminal medulla.

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Manfred Schmidt The central olfactory pathway of the decapod brain is organized into two stages corresponding to the deutocerebral antennular lobes and protocerebral mushroom bodies of the insect brain (Strausfeld et al. 2009). The first stage is constituted by the olfactory deutocerebrum typically containing two prominent paired neuropils, the olfactory lobe (OL) and the accessory lobe (AL) (Fig. 7.5A). Like the insect antennal lobe, the OL receives sensory input from ORNs and is organized into glomeruli, small regions of particularly dense neuropils that are hallmark structures of the first synaptic relay in the central olfactory pathway across higher Bilateria (Sandeman et al. 1992, Schachtner et al. 2005). An AL is present only in Reptantia, and it receives indirect sensory input of various modalities, including prominent input from the OL. The somata of the neurons constituting the olfactory deutocerebrum are segregated into two distinct paired soma clusters (Figs. 7.4A, 7.5A) (Schachtner et al. 2005). The medial soma cluster (MC) contains somata of local interneurons (LNs) with arbors restricted to deutocerebral neuropils, and the lateral soma cluster (LC) contains somata of projections neurons (PNs), which have an axon ascending to the lateral protocerebrum and are differentiated into OL-PNs and AL-PNs in species with large ALs. The lateral protocerebrum represents the second stage of the central olfactory pathway, and it contains LNs whose somata form the hemiellipsoid body cluster (HBC) (Schmidt 1997, Schmidt and Harzsch 1999, Hansen and Schmidt 2001, 2004, Sullivan and Beltz 2005a). Adult Neurogenesis in the Central Olfactory Pathway of the Brain Neuron Counts in the Olfactory Deutocerebrum Counts of neurons in the olfactory deutocerebrum of differently sized juvenile and adult shore crabs Carcinus maenas and crayfish C.  destructor provided direct evidence for a growth-related increase in neuron number (Fig. 7.4D) (Schmidt 1997, Sandeman et al. 1998). In both species, the number of PNs increases substantially and linearly with body size. In C.  maenas, the number of PNs per hemibrain increases from ~12,000 to 24,000, which amounts to a mean addition of at least eight PNs per day in each LC, given that the postlarval life span of C. maenas is approximately four years. In C. destructor, the total number of PNs increases from ~60,000 to 180,000 and the total number of LNs increases from ~30,000 to 50,000. Identification of Proliferation Zones and Deutocerebral “Neurogenic Complexes” To determine whether the continuous increase in neuron number in the olfactory deutocerebrum is based on neurogenesis, in vivo labeling with BrdU was used in conjunction with other microscopical analyses. These experiments, ongoing for the past 15 years mainly in C. maenas, C. destructor, P. argus, and P. clarkii, led to the identification of deutocerebral “neurogenic complexes” (Song et al. 2009) containing the proliferating cells that give rise to new neurons in the olfactory deutocerebrum. A neurogenic complex consists of two compartments connected by a strand- or duct-like structure: (1) a proliferation zone (PZ) within a soma cluster of the olfactory deutocerebrum containing a small group of GMC-like neuronal progenitor cells (GMC-like NPCs), and (2) a putative neurogenic niche (NN) containing one or few primary neuronal progenitor cells that give rise to the GMC-like NPCs in the PZ. Proliferation zones containing groups of BrdU+ cells also occur in soma clusters of the eyestalk ganglia but not in other parts of the CNS (Schmidt 1997). In the eyestalk ganglia of brachyuran and anomuran crabs, but not in other decapods, PZs are present in the HBC (Fig. 7.4C) (Schmidt 1997, Schmidt and Harzsch 1999, Sullivan and Beltz 2005a). Proliferation Zones in Soma Clusters of the Olfactory Deutocerebrum In vivo BrdU labeling with short survival times (6–24 h) led to the identification of groups of BrdU+ cells in or adjacent to the soma clusters of the olfactory deutocerebrum in all decapod



Adult Neurogenesis in Crustaceans

species tested (shrimp:  Sicyonia brevirostris; spiny lobster: P.  argus; clawed lobster:  H.  americanus; crayfish:  C.  destructor, P.  clarkii; anomuran crab:  Pagurus bernhardus; brachyuran crabs: C. maenas, Cancer pagurus, Libinia emarginata) (Schmidt 1997, 2001, 2007a, Sandeman et  al. 1998, Harzsch et  al. 1999, Schmidt and Harzsch 1999, Hansen and Schmidt 2001, 2004, Sullivan and Beltz 2005a, 2005b, Sullivan et al. 2007a, 2007b, Song et al. 2009, Zhang et al. 2009). There is only one group of BrdU+ cells per soma cluster in a specific and invariant position, and the number of BrdU+ cells is small (~20–100) compared to the number of neuronal somata (~10,000–200,000) within the cluster (Figs. 7.4B, 7.5B). Among the group of BrdU+ cells, some cells are in mitosis (Figs. 7.5C, 7.6C, 7.7A), demonstrating that the BrdU+ cells are proliferating and that the areas containing them are PZs. In all tested species, a PZ occurs in the LC, but in the MC, a PZ is present in some species (P. argus, P. interruptus, C. destructor, P. clarkii, L. emarginata) but not in others (S. brevirostris, C. maenas, C. pagurus, P. bernhardus), or it is less well demarcated (H. americanus). Proliferation zones are morphologically distinct from the remainder of the soma clusters in which they reside (Schmidt 1997, 2001, 2007a, 2007b, Schmidt and Derby 2011): (1) The cells in the PZ are significantly smaller and more densely packed than neuronal somata, and they have smaller and more irregularly shaped nuclei (Fig. 7.5C). (2) In P. argus, diverse neuronal markers label neuronal somata in the MC and LC but not the cells in the PZs (Figs. 7.5D, 7.6A,B), demonstrating that the cells in the PZs do not have a neuronal phenotype. (3) Within the PZ of the LC of P. argus, three types of cells (type-A, type-B, type-C cells) with distinct ultrastructure can be distinguished, and all of them differ from mature neuronal somata in shape, size, and nuclear:cytoplasmic ratio (Fig. 7.5E). (4) Neuronal somata are completely separated from each other by interspersed processes of multipolar cell body glia, but type-A and type-B cells are in direct contact with each other and type-C cells are only partially surrounded by glial processes. This parallels the situation in the mushroom bodies of adult crickets, where new Kenyon cells are generated by centrally located NPCs that are not ensheathed by glia (Mashaly et al. 2008). Fate of Proliferating Cells: Terminal Symmetrical Division, Neuronal Maturation, and Apoptosis The fate of the proliferating cells in the PZs was elucidated through BrdU pulse-chase experiments with long survival times (Schmidt 1997, 2001, 2007a, Harzsch et  al. 1999, Hansen and Schmidt 2001, Sullivan and Beltz 2005a, 2005b). These studies revealed the following: (1) Within weeks of labeling, BrdU+ cells leave the PZ and translocate to more peripheral positions within the soma cluster where the somata of mature neurons reside (Figs. 7.6C, 7.7A). (2) Even after very long survival times (up to 14  months in P.  argus), numerous BrdU+ cells are present in peripheral regions of the soma clusters, demonstrating that the cells generated in the PZs survive for a very long time. (3) The number of BrdU+ cells almost exactly doubles between short (14 h in P. argus, 12 h in C. maenas) and much longer survival times (14 days in P. argus, 1 month in C. maenas) indicating that the BrdU+ cells, like GMCs, divide only once (Fig. 7.4E). The morphology of cells in the PZ that are in ana- or telophase of mitosis suggests that as in GMCs this cell division is symmetrical (Figs. 7.6C, 7.7A) (Schmidt 1997, 2001, 2007a, Schmidt and Derby 2011). In C. maenas, the number of BrdU+ cells does not systematically change between 12 and 120 h survival time suggesting that their division proceeds very slowly and likely lasts more than five days (Fig. 7.4E). (4) After survival times of > 1 month, BrdU+ cells can be double-labeled by neuronal markers (antibodies against diverse neuropeptides) and have mature neuronal morphology (ascending axon, arborizations in OL or AL, primary neurite) (Figs. 7.6D–F, 7.7B,C). Thus, the daughter cells generated by the symmetrical division of the BrdU+ cells in the PZ mature into neurons within months. In the LC of C. destructor, new AL-PNs and OL-PNs are produced. In the MC of P. argus, several types of LNs expressing different neuropeptides but

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Fig. 7.6. Cellular organization of neurogenic complexes in the olfactory deutocerebrum of adult decapods. Confocal micrographs. Please see color version of this figure in center plate. (A–C) Proliferation zones. (A and B) MC of Panulirus argus triple labeled with the nuclear marker Hoechst (blue), anti-FMRFamide (FMRF, green), and anti-Substance P (SP, red). The cells composing the proliferation zone (PZ) are FMRF-negative and SP-negative and hence nonneuronal. (C) Labeling with the M-phase marker antipH3 (pH3, green) shows numerous cells in mitosis within the PZ of the LC. All BrdU+ cells (red) labeled by a previous BrdU injection (1 month earlier), have left the PZ. (D–F) Neuronal maturation. (D and E) P. argus. (D) Seven months after BrdU injection, all BrdU+ cells (red) have left the proliferation zone (PZ) of the MC and some of them (inset: arrowhead) are double-labeled by anti-SIFamide (SIF, green) indicating neuronal maturation. Most SIF+ cells are BrdU-negative (inset: asterisk). (E) Six months after BrdU injection, a cell with a BrdU+ nucleus is double labeled by anti-Substance P and has the morphology of a mature neuron as indicated by the presence of a Substance P+ primary neurite (arrowheads). (F) Cherax destructor. Fills of OL-PNs with a red fluorescent dextran (Dex-OL, red) and of AL-PNs with a green fluorescent dextran (Dex-AL, green) several months after exposure to BrdU. Many BrdU+ cells in the LC are double labeled by either dextran (arrowheads: BrdU+/Dex-AL+; double arrowheads: BrdU+/Dex-OL+) demonstrating that newly generated cells mature into AL- and OL-PNs. (G–J) Adult neuroblasts and neurogenic niches. (G and H) P. argus. (G) Adult neuroblast (aNB) in the MC labeled by BrdU (red) after multiple injections over 2 days and by anti-pH3 (pH3, green). Fibrous material surrounding the neurogenic niche (NN) and the duct (D) connecting it with the PZ is nonspecifically labeled by anti-pH3. (H) Labeling of aNB by a single BrdU (red) injection at daytime followed by a survival time of 6 hours. The (Continued)



Adult Neurogenesis in Crustaceans

with unknown morphology are produced. There is no indication that any BrdU+ cells arising from the PZs assume a nonneuronal phenotype. Demonstration of the continuous generation of new neurons raises the question of whether mature neurons are also continuously lost by apoptosis. Initially, darkly stained nuclei in the LC of H. americanus were identified as pyknotic nuclei, and together with results of TUNEL labeling this was interpreted as evidence for widespread apoptosis of mature neurons (Harzsch et al. 1999). Subsequently, similar nuclei in the LC and MC of P. argus were identified as nuclei of cell body glia (Schmidt 2007a, Schmidt and Derby 2011), and labeling with nuclear markers showed that some pyknotic nuclei occur in the vicinity of the PZs but not in the periphery of soma clusters (Schmidt 2001, 2007a). This indicates that some newly generated cells undergo apoptosis as they begin to differentiate but that there is no continuous turnover or replacement of neurons. Modulation of Proliferation by Intrinsic and Extrinsic Factors Proliferation in the PZs of the central olfactory pathway is modulated by internal and external factors. Proliferation was quantified by determining the number of BrdU+ cells in PZs, but this does not provide a measure of the rate of neurogenesis. This is because the number of neurons generated per time interval depends not only on the number of proliferating NPCs but also on the duration of their cell cycle and on the percentage of cells surviving and maturing into neurons (Fig. 7.1B). Internal factors that modulate proliferation are sensory input by ORNs, age, an endogenous circannual rhythm, and serotonin (Hansen and Schmidt 2001, 2004, Sandeman et al. 2009). An external factor that modulates proliferation is environmental richness (Hansen and Schmidt 2004). In C. maenas, unilateral elimination of ORNs (by antennule ablation) reduced the number of BrdU+ cells in the PZs of LC and HBC, and the nature of the effect depended on the temporal order of ablation and BrdU injection (Hansen and Schmidt 2001). When amputation preceded BrdU injection, the number of BrdU+ cells in the PZs of LC and HBC was significantly reduced in both hemibrains compared to controls, indicating that ORN input has a global effect on the replenishment of GMC-like NPCs in the PZs. When BrdU injection preceded amputation, the number of postmitotic BrdU+ cells in the PZs of LC and HBC was significantly lower on the ablated compared to the intact side, indicating that ORN input also inf luences the cell division of the GMC-like NPCs and/ or their survival. In C. maenas, age, season, and captivity modulate proliferation in the PZs of LC and HBC (Fig. 7.4F) (Hansen and Schmidt 2004). With increasing age, the number of

Fig. 7.6. (Continued) aNB is located at the apex of the neurogenic niche (NN). The NN and the duct (D) connecting it with the adjacent PZ are surrounded by fibrous material labeled by anti-pH3 (pH3, green). (I and J) Procambarus clarkii. (I) Labeling with the nuclear marker propidium iodide (PI, blue) reveals that the neurogenic niche consists of two subdivisions (NNS) meeting at the nucleus-free center (arrowhead). The nucleus-free center is labeled by fluorescent dextran (Dex, red) perfused into the cerebral artery and it is surrounded by a layer labeled by anti-Elav (green); arrow (vasculature). (J) Intracellular labeling of NFCs (asterisks) with Lucifer Yellow (LY, green) reveals that they are bipolar extending a short process (arrowheads) to the center of the NN (arrow) and a long process (double arrowheads) toward the PZ. Counter-labeling with anti-GS (GS, blue) and propidium iodide (PI, red). Scale bars: A, C, D = 100 µm; B, F, G–I = 50 µm; D inset, E = 10 µm; J = 20 µm. A, B, E modified from Schmidt 2001 with permission from John Wiley and Sons; D modified from Schmidt 2007b with permission from Oxford University Press; F modified from Sullivan and Beltz 2005b with permission from John Wiley and Sons; G, H modified from Schmidt and Derby 2011 with permission from John Wiley and Sons; I, J modified from Sullivan et al. 2007a with permission from John Wiley and Sons.

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Fig. 7.7. Neuronal maturation in adult Panulirus argus. (A) At a survival time of 1 month after BrdU injection, all BrdU+ cells (light gray) have left the PZ of the LC delineated by cells in mitosis labeled by the M-phase marker anti-pH3 (pH3, white); scale bar = 100 µm. (B and C) At a survival time of 6 months after BrdU injection, a cell in the MC is double-labeled by anti-BrdU and anti-Substance P (SP). Presence of a SP+ primary neurite (arrowheads) demonstrates mature neuronal morphology; scale bar = 10 µm (modified from Schmidt 2001, with permission from John Wiley and Sons).

BrdU+ cells significantly decreased in the LC but remained almost constant in the HBC. In both soma clusters, the number of BrdU+ cells showed seasonal variations with two peaks (spring and late summer) in the LC and one peak (early summer) in the HBC. Keeping the animals in captivity caused a global decrease in the number of BrdU+ cells in the LC and HBC but did not eliminate the seasonal changes. These data suggest that proliferation in the LC and HBC is regulated independently and that an endogenous circannual rhythm contributes to this regulation. In C. destructor, proliferation in the LC continues in an excised and perfused head preparation, albeit with a lower number of BrdU+ cells in the PZ of the LC than in controls (Sandeman et al. 2009). Serotonin, either systemically applied via the brain perfusion or released from serotonergic neurons after electrical stimulation of the olfactory deutocerebrum, caused an increase in the number of BrdU+ cells in the LC back to the control level. Adult Neuroblasts After the initial identification of GMC-like NPCs in PZs of the LC and MC, several years passed until their origin began to be elucidated. This delay was mainly because in early experiments on C. maenas and P. argus, single BrdU injections were given in the evening, and this only labeled cells inside of PZs, where none are retained after longer survival times (Schmidt 1997, 2001). In P. argus, multiple BrdU injections over ~2 days led to the discovery of one additional BrdU+ cell that is located outside of each PZ within a dense clump of small cells that is connected by a short duct to the respective PZ and likely represents a neurogenic niche (Figs. 7.6G, 7.8A) (Schmidt 2002, 2007a; see below). Based on having a significantly larger nucleus than GMC-like NPCs and mature neurons, these extra BrdU+ cells were identified as adult NBs (aNBs), since the distinguishing feature of embryonic and larval NBs of Tetraconata is to be larger and to have a larger nucleus than GMCs and neurons (Schmidt 2007a, 2007b, Egger et al. 2008). Subsequent experiments showed that the aNBs of P. argus are also labeled by single BrdU injections followed by short survival times (4–8 h) if the BrdU injections are given at daytime (Figs. 7.6H,



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Fig. 7.8. Adult neuroblasts and neurogenic niches in Panulirus argus. (A) Methylene blue stained semithin section through the MC reveals that the PZ is connected via a duct, D, to the adjacent NN containing a large aNB in metaphase of mitosis; N (soma of neuron); scale bar = 10 µm (modified from Schmidt 2007a, with permission from John Wiley and Sons). (B) Triple labeling by anti-BrdU (single BrdU injection at daytime; survival time: 6 hours), anti-pH3, and Hoechst. The large BrdU+ aNB nucleus is located at the apex of the NN. The NN and the duct, D, connecting it with the adjacent PZ are surrounded by fibrous material labeled by anti-pH3; N (nucleus of neuron); scale bar = 50 µm (modified from Schmidt and Derby 2011, with permission from John Wiley and Sons). (C) TEM micrograph of the NN in the LC. Small somata of niche-forming cells (NFC) form a dense cortex around the nucleus-free center that contains the bulbous foot of the aNB; scale bar = 10 µm; inset: highlighted interface between bulbous foot of aNB (light gray) and inner NFC processes (dark gray); scale bar = 1 µm (modified from Schmidt and Derby 2011, with permission from John Wiley and Sons). (D) Reconstruction of the NN in the LC (modified from Schmidt and Derby 2011, with permission from John Wiley and Sons). Abbreviations: A, arteriole; aNB, adult neuroblast; G, cell body glia; NPC, neural progenitor cell; NFC, niche-forming cell; PVC, perivascular cell; bulbous foot of aNB (asterisk).

7.8B) (Schmidt and Derby 2011), suggesting that the initial failure to label aNBs with single BrdU injections was due to the wrong time of day of the injections. Thus, the aNBs of P. argus are not largely quiescent but divide frequently and their cell divisions appear to be under circadian control. The analysis of the ultrastructure and the mitotic plane of the aNBs of P. argus provided evidence that they undergo asymmetric, self-renewing cell divisions (Fig. 7.8C, D) (Schmidt and Derby 2011). The aNBs of P. argus have a unique, hourglass-like shape with two domains connected by a thin cytoplasmic bridge. The outer domain containing the large nucleus is located at the PZ-facing pole of the NN, where the duct connecting it with the PZ arises, and the inner domain forms a bulbous foot in the center of the NN. The mitotic plane of the aNB nucleus is perpendicular to the long axis of the hourglass. This suggests that in each cell division an outer daughter cell is budded off from the outer aNB domain into the duct, whereas the inner daughter cell retains the hourglass-like shape and represents the self-renewed aNB. Spindle-shaped cells within the duct, some of which are BrdU+, likely represent outer daughter cells migrating toward the PZ to replenish the pool of GMC-like NPCs (demonstrated in P. clarkii for homologous cells: Sullivan et al. 2007a, Zhang et al. 2009). Putative aNBs with significantly larger nuclei than GMC-like NPCs were also identified in adult P. clarkii by single BrdU injections followed by short survival times (Fig. 7.9C) (Song et al. 2009). As in P. argus, one putative aNB is associated with each PZ of MC and LC, the aNBs are located within NNs connected to the PZs, and the orientation of their mitotic plane is perpendicular to the NN-PZ axis (Song et al. 2009, Zhang et al. 2009). This suggests that in P. clarkii, as in P. argus, the putative aNBs bud off daughter cells toward the associated PZ through repeated asymmetric, self-renewing cell divisions. An alternative model of the cellular events leading to

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Manfred Schmidt replenishment of the pool of GMC-like NPCs has been proposed (Zhang et al. 2009, Sandeman et al. 2011; see below). Neurogenic Niches Neurogenic niches were originally described as “deutocerebral organs” in ~20 species of decapods including representatives of most major infraorders, but without connection to neurogenesis (Bazin and Demeuzy 1968, Bazin 1969, 1970a, 1970b). They were rediscovered independently in P. argus and P. clarkii with in vivo BrdU labeling experiments, and initially various terms (“sheath of glial cells,” “glial soma cluster,” “clump of cells”) were used for them (Schmidt 2002, 2007a, Song et al. 2007, 2009, Sullivan et al. 2007a, Zhang et al. 2009). In all species, the NNs have a unique morphology featuring a densely packed cortex of small, darkly stained cells (niche-forming cells  =  NFCs) surrounding a nucleus-free center (Figs. 7.8B–D, 7.9E inset). Across decapods, the NNs are invariably associated with the soma clusters of the olfactory deutocerebrum, but there are substantial interspecific differences in their location. The NNs are located within the soma clusters in spiny lobsters, slipper lobsters, and brachyuran crabs, between LC and MC at the ventral surface of the brain in crayfish and anomuran crabs, or medially between the roots of the antenna II nerves and attached to a lamellated ovoid body at the outside of the brain in clawed lobsters. In crayfish, there is one NN per hemibrain that is composed of two subdivisions, one connected with the PZ in the LC and the other one with the PZ in the MC via long strands of fibrous tissue (Fig. 7.9A,E inset). The linkage of the NNs to the PZs in the soma clusters of the olfactory deutocerebrum and their association with aNBs that likely act as self-renewing NSCs, led to the hypothesis that they represent neural stem cell niches and are indispensable components of adult neurogenesis across

Fig. 7.9. Putative adult neuroblasts and neurogenic niches in Procambarus clarkii. (A)  Neurogenic complex of one hemibrain labeled by anti-glutamine synthetase (white); LPZ (PZ of LC), MPZ (PZ of MC); scale bar = 100 µm (modified from Sullivan et al. 2007a, with permission from John Wiley and Sons). (B) NFCs (asterisks) injected with Lucifer Yellow (white) are bipolar with a short process (arrowheads) projecting to the center of the NN (arrow) and a long process (double arrowheads) projecting to the associated PZ; scale bar = 20 µm (modified from Sullivan et al. 2007a, with permission from John Wiley and Sons). (C and D) BrdU pulse-chase experiments with 6 h survival time (modified from Song et al. 2009, with permission from Elsevier Limited). (C) Typically one large putative aNB (arrowhead) in one subdivision (S) of the NN is BrdU+ (white); Hoechst-labeled (light gray) elongated nuclei of NFCs (arrow); nucleolus of aNB (double arrowhead). (D) Rarely NFCs with flat elongated nuclei (visualized by Hoechst, light gray) are BrdU+ (white, arrows); scale bar = 10 µm (also applies to C). (E) TEM micrograph of the nucleus-free center of the NN. Inner processes of NFCs surround an extracellular space (asterisk) filled with cuticle-like material. The NFC processes carry apical microvilli (MV) and are interconnected subapically by adherens junctions (arrows); scale bar = 1 µm. Inset: semithin section stained with Methylene blue reveals that the NN consists of two subdivisions (S) meeting at the nucleus-free center (asterisk); scale bar = 50 µm.

Fig. 7.10. Models of adult brain neurogenesis. (A) Unified model of adult neurogenesis in the olfactory deutocerebrum of decapods (modified from Schmidt and Derby 2011, with permission from John Wiley and Sons). In each neuronal soma cluster, neurogenesis is based on one NSC, a large aNB embedded in a neurogenic niche composed of niche-forming cells (NFC). Through rapid asymmetric divisions, the aNB self-renews and produces differentiating daughter cells. Within a duct formed by NFC processes, the daughter cells migrate to the PZ, where they replenish the pool of GMC-like NPCs (NPC). GMC-like NPCs divide once symmetrically and produce two immature neurons (iN). Migrating away from the PZ, most iNs become differentiated neurons (dN) within months but some die by apoptosis. Daughter cells generated by the aNB may undergo one or more fast rounds of cell divisions and thus represent intermediate progenitor cells (IPC?). NFCs may occasionally replace the aNB (dotted line with?). (B) Hypothetical embryonic origin of the neurogenic niche. (1 and 2) Invagination of a patch of neuroectodermal cells containing a globose NB, producing embryonic cuticle apically, and being subapically connected by adherens junctions (dark gray boxes). (3) Closure of the invaginated patch leads to a configuration as in the NN of crayfish. The nucleus-free center is filled by cuticle-like material produced by inner NFC processes subapically connected by adherens junctions. (4) Expansion of the apical pole of the aNB leads to a configuration as in the NN of P. argus; the aNB buds off GMC-like NPCs at its basal pole. (C) Comparative models of embryonic and adult neurogenesis in mammals, crustaceans, and insects (from Schmidt and Derby 2011, with permission from John Wiley and Sons). Mammals: Ciliated (arrowhead) radial glial cells serve as NSCs in the embryonic CNS and generate neurons either directly or through intermediate progenitor cells (IPC?). Radial glial cells give rise to SVZ astrocytes (type B cells) in the adult brain that are also ciliated (arrowhead) and act as slowly cycling NSCs. Type B cells are associated with complex neural stem cell niches. Crustaceans: Neurogenesis in the embryonic and larval CNS is based on large globose NBs that self-renew and generate GMCs through asymmetric divisions. Each GMC divides symmetrically once and generates two immature neurons. Adult neurogenesis in the olfactory midbrain is based on few large aNBs that cycle rapidly undergoing self-renewing asymmetric divisions. Ultimately they give rise to GMC-like NPCs, each generating two immature neurons (left lineage). Daughter cells produced by the aNBs may act as intermediate progenitor cells (IPC; right lineage). Each aNB is closely associated with a discrete neurogenic niche. Insects: Neurogenesis in embryonic and larval CNS is based on large globose NBs. Most NBs give rise to neurons through the canonical lineage as in crustaceans (left lineage). Some NB lineages contain intermediate progenitor cells (IPC) that self-renew and generate GMCs (right lineage). Adult neurogenesis in the mushroom body is maintained by canonical NBs that are not associated with a stem cell niche.

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Manfred Schmidt decapods (Schmidt 2007a, 2007b, Sullivan et al. 2007a, 2007b, Song et al. 2009, Sandeman et al. 2011). Stem cell niches are specialized cellular microenvironments in which adult stem cells in various tissues reside, and they provide regulatory input and nutritional support to the adult stem cells and shield them from harmful factors (e.g., Morrison and Spradling 2008). The NSCs supporting adult neurogenesis in mammals are associated with stem cell niches composed of glia, vascular cells, and extracellular matrix structures that are dispersed throughout the germinal layers (e.g., Shen et al. 2008, Tavazoie et al. 2008). That the NNs represent neural stem cell niches is supported by more detailed analyses of their cytoarchitecture in P. argus and P. clarkii (Schmidt 2007a, Song et al. 2007, 2009, Sullivan et al. 2007a, Zhang et al. 2009, Schmidt and Derby 2011). The NNs consist of a structurally homogeneous and numerically constant population of NFCs (P. clarkii: ~300; P. argus: ~120 in LC, ~85 in MC) that have a distinct bipolar morphology with a short process projecting inward and a long process projecting outward to the associated PZ (Figs. 7.6J, 7.8D, 7.9B). In P. argus, the inner processes of the NFCs completely cover the bulbous foot of the aNB that fills the nucleus-free center of the NN and the outer NFC processes envelope the outer domain of the aNB and fill the interior of the duct (Fig. 7.8C,D). Thus, the NFCs are the exclusive constituents of the aNB microenvironment. In crayfish, labeling of the nucleus-free center after perfusion of fluorescent tracers into the cerebral artery led to the conclusion that it represents a vascular cavity (Fig. 7.6I) (Sullivan et al. 2007a, Sandeman et  al. 2009). This, however, is in conflict with data from transmission electron microscopy (TEM) that fail to show a direct connection of the nucleus-free center with surrounding arterioles and demonstrate that it is filled by cuticle-like material (Fig. 7.9E) (Sandeman et al. 2011). Prerequisite for understanding how the NNs function is knowing the cellular identity of the NFCs. Niche-forming cells are likely homologous across decapods and therefore should have the same cellular identity in all species, but this issue is still controversial. In P. clarkii, the NFCs were identified as glial cells (Sullivan et al. 2007a) based on being labeled by antiglutamine synthetase (anti-GS) (Figs. 7.6J, 7.9A), a selective marker for astrocytes in mammals (Norenberg 1979) and for neuropil glia in decapod brains (Linser et al. 1997, Sullivan et al. 2007a, Harzsch and Hansson 2008, Schmidt and Derby 2011). In P. argus and the anomuran crab Coenobita clypeatus, anti-GS selectively labeled neuropil glia but failed to label NFCs, which does not support the identification of NFCs as glial cells (Harzsch and Hansson 2008, Schmidt and Derby 2011). The distinct bipolar morphology of NFCs is further evidence against their glial identity, since crustacean glial cells are predominantly uni- or multipolar (e.g., Cuadras and Marti-Subirana 1987). In P. argus, TEM and light microscopy using markers for different cell types revealed that the NFCs do not correspond to any cell type in the olfactory deutocerebrum, including neurons, several types of glia, perivascular cells, and hemocytes (Schmidt and Derby 2011). Thus, NFCs appear to represent an unidentified cell type and Schmidt and Derby (2011) proposed that they correspond to embryonic neuroepithelial cells, which also have a distinct bipolar morphology and in basal insects remain associated with the delaminated embryonic NBs as sheath cells (Doe and Goodman 1985). Development of Neurogenic Complexes in Juveniles The development of the neurogenic complexes was traced in P. clarkii from hatchlings to adults (Song et al. 2007, 2009, Zhang et al. 2009). Neurogenic complexes are already identifiable in the first postembryonic stage (POI) and hence must originate during embryogenesis. In subsequent development, the neurogenic complexes undergo several major changes: (1) The number of proliferating cells declines dramatically in all compartments of the neurogenic complexes until in adults only very few BrdU+ cells are present in the PZ of the MC and in the NNs. (2) In hatchlings, the PZs of MC and LC and the NN between them are very close together and become ever more separated due to growth of the brain. (3) The ever-increasing distance between the



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PZs and the NN is compensated by the simultaneous elongation of the strands connecting each subdivision of the NN with the associated PZ. (4) The NN increases substantially in length (50 µm in POI/~250 µm in adult) and in the number of NFCs (< 100 in POI/≥ 300 in adult). The increase in the number of NFCs appears to be based on local NFC proliferation (Zhang et al. 2009), which even in adults occurs occasionally (Fig. 7.9D). Several studies have reported the modulation of proliferation in the neurogenic complexes of early juvenile crayfish and clawed lobsters by diverse internal and external factors, including environmental richness (Sandeman and Sandeman 2000, Ayub et al. 2011), social status (Song et al. 2007), circadian time (Goergen et al. 2002), nitric oxide (Benton et al. 2007), omega-3 fatty acids (Beltz et al. 2007), and serotonin (Benton et al. 2008, Zhang et al. 2011). Models of Adult Neurogenesis in the Olfactory Deutocerebrum Two models of adult neurogenesis in the olfactory deutocerebrum have been developed. Model A (Sullivan et al. 2007a, 2007b, Zhang et al. 2009, Sandeman et al. 2011) holds that NFCs have glial cell identity and act as “first-generation neuronal precursor cells.” The NFCs divide symmetrically and do not self-renew and thus do not represent NSCs. Both daughter cells generated by a NFC division (“second-generation precursors”) migrate away from the NN to the associated PZ. In the PZ, these cells divide at least once more and their progeny (“third and subsequent generations”) differentiate into neurons. Since the NFCs do not self-renew, their population needs to be continuously replenished, and it is proposed that this is through transdifferentiation of hemocytes reaching the NN via its “vascular cavity.” The assumed glial identity of the NFCs and their association with a “vascular cavity” leads to the conclusion that adult neurogenesis in decapods is similar to adult neurogenesis in the mammalian brain, which is maintained by astrocytic NSCs associated with stem cells niches composed of glial and vascular elements (Fig. 7.10C) (Kriegstein and Alvarez-Buylla 2009). Model B (Schmidt 2007a, 2007b, Song et al. 2009, Schmidt and Derby 2011) holds that one large aNB associated with each PZ acts as an NSC. Through repeated and rapid asymmetric divisions, the aNB self-renews and buds off a daughter cell at its PZ-facing pole. These daughter cells migrate to the associated PZ, where they give rise to GMC-like NPCs. This process probably involves early, as yet unidentified, proliferation events in which the aNB daughters act as transit amplifying intermediate progenitor cells as in some NB lineages of insects (e.g., Boyan and Reichert 2011). The GMC-like NPCs undergo one slow terminal symmetrical division, in which two immature neurons are generated that migrate away from the PZ. Some of them die early through apoptosis, while most survive and differentiate into neurons within months. Each aNB is closely associated with an NN supporting the long-term survival and proliferative activity of the aNB. The NN is composed of bipolar NFCs that correspond to embryonic neuroepithelial cells. Since basal developmental processes are generally conserved in closely related species, it is unlikely that adult neurogenesis in the olfactory deutocerebrum proceeds in fundamentally different ways in crayfish and spiny lobsters (Sandeman et al. 2011). A unified view of adult neurogenesis reconciling the two conflicting models (Fig. 7.10A) is based on the new interpretation of the NNs as being derived from patches of embryonic neuroepithelium through a process of invagination (Fig. 7.10B) as in imaginal disks and optic lobe primordia of holometabolous insects and groups of neuronal precursors in spider embryos (Green et al. 1993, McClure and Schubiger 2007, Stollewerk 2008). If so, the NFCs could behave like neuroepithelial or imaginal disk cells, which multiply through symmetrical divisions and—as a prelude to neurogenesis—single out cells that enlarge, delaminate, and switch to a NB fate (Egger et al. 2007). In this view, NFCs could expand in number through symmetrical divisions and—at least occasionally—serve as “first-generation neuronal precursor cells” as posited by Model A. However, this would be achieved by a selection process in which one NFC is specified as an aNB, which then acts as self-renewing NSC as posited by Model B. Thus, the unified model proposes that

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Manfred Schmidt adult neurogenesis in the olfactory deutocerebrum of decapods is essentially identical to embryonic and larval neurogenesis in Tetraconata, in being based on embryonic neuroepithelial cells (NFCs) and NBs derived from them (aNBs) (Fig. 7.10C). A key component of adult neurogenesis not present in embryonic and larval neurogenesis is that the aNBs remain permanently associated with the NFCs forming an NN that preserves an embryonic microenvironment. Adult Neurogenesis in Other Parts of the Central Nervous System Outside the central olfactory pathway, the only regions of the CNS of adult decapods in which PZs have been identified are the optic lobes. In C. maenas and C. destructor, PZs occur in soma clusters in the optic lobes, and they are typically located at the lateral edge of the optic neuropils (Schmidt 1997, Sullivan and Beltz 2005a). Some BrdU-pulse chase experiments and double-labeling with neuronal markers demonstrated that BrdU+ cells originating from these PZs mature into neurons (Sullivan et al. 2005a).

CONSTITUTIVE NEUROGENESIS IN THE ADULT BRAIN: COMPARISON BETWEEN TAXA As detailed in the Introduction, constitutive neurogenesis in the adult brain is common among three bilaterian taxa: vertebrates, insects, and decapods. Comparing how neurogenesis is maintained in these taxa is key to understanding the evolutionary history, cellular basis, and functional significance of this profound and long-term neuronal plasticity (Fig.  7.10C). According to the unified model, adult neurogenesis in decapods has obvious commonalities with adult neurogenesis in insects with respect to the identity of the NSCs and the cell lineages they produce. In both taxa, the NSCs maintaining adult neurogenesis are few, large aNBs of invariant location within or close to the neuronal soma clusters in which new neurons are generated (Cayre et al. 2007). Through successive asymmetric divisions, the aNBs ultimately give rise to NPCs (GMCs, GMC-like NPCs) which, through one terminal symmetrical division, generate two neurons. To produce the high number of GMC-like NPCs associated with each aNB in decapod adult neurogenesis, at least some daughter cells generated by the aNBs should act as transit amplifying intermediate progenitor cells (Schmidt and Derby 2011), as has been described for some embryonic and larval NB lineages of insects (e.g., Boyan and Reichert 2011). However, transit amplifying intermediate progenitor cells do not occur in the NB lineages producing Kenyon cells, the only neurons generated in adults (Cayre et  al. 2007). Another substantial difference between both taxa is that in decapods but not in insects, NBs are associated with neural stem cell niches, since in the Kenyon cell layer of adult crickets where adult neurogenesis occurs, no particular association of NPCs with glial cells is evident (Mashaly et  al. 2008). Presumably, the presence or absence of a stem cell niche associated with aNBs reflects the enormous difference in the duration of adult neurogenesis between decapods and insects. In decapods, adult neurogenesis persists throughout adult life lasting from years to decades (see above), whereas the adult life span of insects is in the range of months and adult neurogenesis typically ceases within the first weeks of adulthood (e.g., Zhao et al. 2008). Thus, adult neurogenesis in insects is a transient continuation of larval/pupal neurogenesis, and it may be enabled by preventing apoptotic death of mushroom body NBs (Siegrist et al. 2010). In contrast, adult neurogenesis in decapods is a lifelong process and appears to require that stem cell niches preserving an embryonic microenvironment support the survival and proliferative potential of aNBs. Adult neurogenesis in decapods and insects differs fundamentally from adult neurogenesis in mammals with respect to NSC identity, number, and proliferation dynamics. In decapods and



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insects, adult neurogenesis is maintained by tetraconate-typical NBs that are large, do not have a differentiated cell phenotype, lack a primary cilium, are few in number, and reside in invariant locations. In contrast, adult neurogenesis in mammals is maintained by NSCs that have an astrocyte phenotype, bear a primary cilium, occur in large number, and are dispersed throughout extensive germinal layers (Kriegstein and Alvarez-Buylla 2009). The aNBs of decapods and insects proliferate actively (Zhao et al. 2008, Song et al. 2009, Schmidt and Derby 2011), whereas NSCs of mammals are largely quiescent (Doetsch et al. 1999). Aside from these substantial differences in the nature of the NSCs, adult neurogenesis in decapods (but not in insects) parallels adult neurogenesis in mammals. First, adult neurogenesis is maintained throughout adult life, which in decapods and mammals can last several years to decades. Second, in both taxa the NSCs are supported by stem cell niches, which likely preserve elements of the embryonic microenvironment and in addition include glial and vascular elements. A generality uniting adult neurogenesis in decapods, insects, and mammals is that in each taxon the NSCs maintaining adult neurogenesis have the same identity as those fueling embryonic (and larval) neurogenesis. Thus, adult neurogenesis can be viewed as an extension of embryonic neurogenesis (Kriegstein and Alvarez-Buylla 2009). Furthermore, the presence of stem cell niches associated with the NSCs fueling adult neurogenesis in decapods and mammals, but not in insects, suggests that neural stem cell niches are a prerequisite for preserving the proliferative activity of NSCs for years or even decades. To understand the mechanisms by which neural stem cell niches achieve this, it will be informative to compare those of decapods and mammals, especially because they are of different cellular composition.

OPEN QUESTIONS AND FUTURE DIRECTIONS Adult neurogenesis in the PNS and CNS is an attractive and active field of crustacean developmental neurobiology with many intriguing directions still unexplored. Some of the open questions that will be the focus of future exploration are outlined below. Is the Generation of New Sensilla throughout Life in Decapods Based on Equivalent Cell Lineages and Regulatory Gene Networks as in Insects? Crustacean sensilla have substantially more receptor neurons and auxiliary cells than insect sensilla, and this is most striking in aesthetascs, which typically house several hundred ORNs. Two alternative scenarios seem plausible:  (1)  all cells of a sensillum originate from one SOP through a common cell lineage containing one or more self-renewing progenitor cells, and (2) the cells of a sensillum arise from an assemblage of nonlineage related ectodermal cells as in olfactory sensilla of insects (e.g., Reddy et al. 1997). How Does the Neurogenic Niche Contribute to the Lifelong Maintenance of Adult Neuroblasts? A general problem in stem cell biology is how the genomic integrity of stem cells is maintained in spite of the accumulation of DNA copy errors through repeated rounds of asymmetric divisions. The common feature of stem cells to be mainly quiescent is interpreted as a mechanism to avoid accumulation of copy errors by minimizing the number of divisions. Since the aNBs of decapods divide actively, there must be other mechanisms in place to maintain their genomic integrity. One plausible mechanism suggested by the unified model of adult neurogenesis is that

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Manfred Schmidt an aNB has only a limited life span, after which it is replaced by a new aNB arising from the pool of associated NFCs (Fig. 7.10A). Where Is the Evolutionary Origin of Adult Neurogenesis in Crustaceans? Adult neurogenesis appears to be present ubiquitously across decapods, but it is unknown whether it also occurs in other crustacean taxa. Assaying representatives of other orders within Malacostraca and beyond should reveal whether adult neurogenesis is an evolutionary old trait or a more recent development within Malacostraca. What Is the Function of Adult Neurogenesis in the Olfactory Pathway? Arguably the most interesting question about adult neurogenesis in the olfactory pathway of decapods is what function it serves. Two different levels are to be considered: (1) Continuous addition of aesthetascs with a fixed complement of ORNs likely causes an increase in odor sensitivity. Thus older adults should have a higher odor sensitivity than younger ones, and this could be tested with behavioral assays based on aesthetasc-specific responses (e.g., Shabani et  al. 2008). (2) The continuous addition of PNs (and LNs) to the olfactory deutocerebrum may provide a means to adapt the processing of olfactory information to ever-changing odor environments (Schmidt 2007b). In this scenario, new interneurons could connect with and thus encode novel combinations of activated OL glomeruli that emerge when novel odors are encountered. Thus, experimentally blocking adult neurogenesis should result in a decreased ability to appropriately respond to novel odors. By providing a meaningful comparison with current efforts to unravel the importance of adult neurogenesis for olfaction in mammals (e.g., Lazarini and Lledo 2011), this research could uncover fundamentally important principles of the role of adult neurogenesis in olfactory information processing.

SUMMARY AND CONCLUSIONS This chapter provides an overview of adult neurogenesis in the PNS and CNS of decapod crustaceans, most of which continue to grow indeterminately as adults. In these species, adult neurogenesis of receptor neurons in the PNS is nearly ubiquitous and linked to the addition of entire sensory units to the exoskeleton, incrementally expanding in each molt. In contrast, adult neurogenesis of neurons in the CNS, present in all species thus far analyzed, is restricted to only two areas, the visual and the olfactory pathways. Adult neurogenesis in the visual pathway seems to be linked to the addition of retinotopic “columns” to the neuropils of the optic lobes associated with the molt-related increase in the number of ommatidia in the compound eyes. In contrast, adult neurogenesis in the central olfactory pathway is a continuous process independent of the addition of new ORNs, and it is based on proliferation in small zones located deep within the clusters of somata of mature neurons. In the olfactory deutocerebrum, which includes the first synaptic relay of the central olfactory pathway, the OL, each PZ is associated with one putative neural stem cell, a large aNB. Each aNB is embedded in a presumptive NN composed of small, bipolar cells of controversial identity (glial vs. ectodermal). Adult neurogenesis in decapods parallels adult neurogenesis in the mammalian brain in being based on the same type of neuronal stem cells as embryonic neurogenesis and in having NNs associated with the neural stem cells. Elucidating further the cellular organization and functional properties of the NNs in decapod crustacean brains



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will contribute to a broader understanding of how life-ong adult neurogenesis is maintained and regulated.

ACKNOWLEDGMENTS Original work has been supported by NIH grant DC00312. I  thank Charles Derby and two anonymous reviewers for many constructive comments, which helped to significantly improve this manuscript.

REFERENCES Abbott, N.J. 1971. The organization of the cerebral ganglion in the shore crab, Carcinus maenas. II. The relation of intracerebral blood vessels to other brain elements. Zeitschrift für Zellforschung 120:401–419. Ayub, N., J.L. Benton, Y. Zhang, and B.S. Beltz. 2011. Environmental enrichment influences neuronal stem cells in the adult crayfish brain. Developmental Neurobiology 71:351–361. Bazin, F. 1969. Étude comparée d’un organe deutocérébral chez les Crustacés Décapodes Reptantia. Comptes Rendus de l’Académie des Sciences. Série D 269:958–961. Bazin, F. 1970a. Étude comparée de l’organe deutocérébral des Macroures Reptantia et des Anomures (Crustacés Décapodes). Archives de Zoologie Expérimentale et Générale 111:245–264. Bazin, F. 1970b. Les organes deutocérébraux chez deux Crustacés Décapodes Macroures Reptantia: Panulirus regius De Brito Capello, Scyllarus arctus (L.). Bulletin de la Societe Zoologique de France 96:87–92. Bazin, F., and N. Demeuzy. 1968. Existence d’organes intracérébraux énigmatiques chez le Crustacé Décapode Carcinus maenas (L.). Comptes Rendus de l’Académie des Sciences. Série D 267:356–358. Beltz, B.S., M.F. Tlusty, J.L. Benton, and D.C. Sandeman. 2007. Omega-3 fatty acids upregulate adult neurogenesis. Neuroscience Letters 415:154–158. Benton, J.L., D.C. Sandeman, and B.S. Beltz. 2007. Nitric oxide in the crustacean brain: regulation of neurogenesis and morphogenesis in the developing olfactory pathway. Developmental Dynamics 236:3047–3060. Benton, J.L., E.M. Goergen, S.C. Rogan, and B.S. Beltz. 2008. Hormonal and synaptic influences of serotonin on adult neurogenesis. General and Comparative Endocrinology 158:183–190. Bernhards, H. 1916. Der Bau des Komplexauges von Astacus fluviatilis (Potamobius astacus L.). Ein Beitrag zur Morphologie der Decapoden. Zeitschrift für wissenschaftliche Zoologie 116:649–707. Boyan, G.S., and H. Reichert. 2011. Mechanisms for complexity in the brain: generating the insect central complex. Trends in Neurosciences 34:247–257. Carlisle, D.B. 1957. On the hormonal inhibition of moulting in decapod Crustacea. II. The terminal anecdysis in crabs. Journal of the Marine Biological Association of the United Kingdom 36:291–307. Cayre, M., S. Scotto-Lomassese, J. Malaterre, C. Strambi, and A. Strambi. 2007. Understanding the regulation and function of adult neurogenesis: contribution from an insect model, the house cricket. Chemical Senses 32:385–395. Chien, H., T. Tadesse, H. Liu, M. Schmidt, W.W. Walthall, P.C. Tai, and C.D. Derby. 2009. Molecular cloning and characterization of homologs of achaete-scute and hairy-enhancer of split in the olfactory organ of the spiny lobster Panulirus argus. Journal of Molecular Neuroscience 39:294–307. Cooper, R.L., and C.K. Govind. 1991. Axon composition of the proprioceptive PD nerve during growth and regeneration of lobster claws. Journal of Experimental Zoology 260:181–193. Cuadras, J., and A. Marti-Subirana. 1987. Glial cells of the crayfish and their relationships with neurons. An ultrastructural study. Journal of Physiology (Paris) 82:196–217. Curtis, M.A., B. Connor, and R.L.M. Faull. 2003. Neurogenesis in the diseased adult human brain. New therapeutic strategies for neurodegenerative diseases. Cell Cycle 2:428–430.

199

200

Manfred Schmidt Derby, C.D., H.S. Cate, P. Steullet, and P.J.H. Harrison. 2003. Comparison of turnover in the olfactory organ of early juvenile stage and adult Caribbean spiny lobsters. Arthropod Structure and Development 31:297–311. Doe, C.Q., and C.S. Goodman. 1985. Early events in insect neurogenesis. I. Development and segmental differences in the pattern of neuronal precursor cells. Developmental Biology 111:193–205. Doetsch, F., I. Caillé, D.A. Lim, J.M. García-Verdugo, and A. Alvarez-Buylla. 1999. Subventricular zone astrocytes are neural stem cells in the adult mammalian brain. Cell 97:703–716. Dohle, W. 1976. Die Bildung und Differenzierung des postnauplialen Keimstreifs von Diastylis rathkei (Crustacea, Cumacea). II. Die Differenzierung und Musterbildung des Ektoderms. Zoomorphologie 84:235–277. Dohle, W. 2001. Are the insects terrestrial crustaceans? A discussion of some new facts and arguments and the proposal of the proper name “Tetraconata” for the monophyletic unit Crustacea + Hexapoda. Annales de la Société Entomologique de France 37:85–103. Duman-Scheel, M., and N.H. Patel. 1999. Analysis of molecular marker expression reveals neuronal homology in distantly related arthropods. Development 126:2327–2334. Egger, B., J.O. Boone, N.R. Stevens, A.H. Brand, and C.Q. Doe. 2007. Regulation of spindle orientation and neural stem cell fate in the Drosophila optic lobe. Neural Development 2:1. Egger, B., J.M. Chell, and A.H. Brand. 2008. Insights into neural stem cell biology from flies. Philosophical Transactions of the Royal Society B—Biological Sciences 363:39–56. Eguchi, E., K. Arikawa, S. Ishibashi, T. Suzuki, and V.B. Meyer-Rochow. 1989. Growth-related biometrical and biochemical studies of the compound eye of the crab, Hemigrapsus sanguineus. Zoological Science 6:241–250. Ekerholm, M., and E. Hallberg. 2002. Development and growth patterns of olfactory sensilla in malacostracan crustaceans. Pages 376–385 in K. Wiese, editor. The crustacean nervous system. Springer-Verlag, Berlin. Farmer, A.S. 1973. Age and growth in Nephrops norvegicus (Decapoda: Nephropidae). Marine Biology 23:315–325. Farris, S.M., and I. Sinakevitch. 2003. Development and evolution of the insect mushroom bodies: towards the understanding of conserved developmental mechanisms in a higher brain center. Arthropod Structure and Development 32:79–101. Field, L.H. 2005. The chordotonal organ: a uniquely invertebrate mechanoreceptor. Pages 61–105 in T.A. Christensen, editor. Methods in insect sensory neuroscience. CRC Press, Boca Raton, London, New York, Washington. Finley, L., and D. Macmillan. 2000. The structure and growth of the statocyst in the Australian crayfish Cherax destructor. Biological Bulletin 199:251–256. Gerberding, M. 1997. Germ band formation and early neurogenesis of Leptodora kindti (Cladocera): first evidence for neuroblasts in the entomostracan crustaceans. Invertebrate Reproduction and Development 32:63–73. Goergen, E.M., L.A. Bagay, K. Rehm, J.L. Benton, and B.S. Beltz. 2002. Circadian control of neurogenesis. Journal of Neurobiology 53:90–95. Govind, C.K., and D.J. Potter. 1987. Development of bilateral asymmetry in sensory innervation to lobster claws. Developmental Brain Research 35:131–139. Green, P., A.Y. Hartenstein, and V. Hartenstein. 1993. The embryonic development of the Drosophila visual system. Cell and Tissue Research 273:583–98. Gross, C.G. 2000. Neurogenesis in the adult brain: death of a dogma. Nature Reviews Neuroscience 1:67–73. Grünert, U., and B.W. Ache. 1988. Ultrastructure of the aesthetasc (olfactory) sensilla of the spiny lobster, Panulirus argus. Cell and Tissue Research 251:95–103. Guse, G.W. 1983. Ultrastructure, development, and moulting of the aesthetascs of Neomysis integer and Idotea baltica (Crustacea, Malacostraca). Zoomorphology 103:121–133. Hafner, G.S., and T.R. Tokarski. 2001. Retinal development in the lobster Homarus americanus— comparison with compound eyes of insects and other crustaceans. Cell and Tissue Research 305:147–158.



Adult Neurogenesis in Crustaceans

Hallberg, E., and M. Skog. 2011. Chemosensory sensilla in crustaceans. Pages 103–121 in T. Breithaupt and M. Thiel, editors. Chemical communication in crustaceans. Springer, New York. Hansen, A., and M. Schmidt. 2001. Neurogenesis in the central olfactory pathway of the adult shore crab Carcinus maenas is controlled by sensory afferents. Journal of Comparative Neurology 441:223–233. Hansen, A., and M. Schmidt. 2004. Influence of season and environment on adult neurogenesis in the central olfactory pathway of the shore crab, Carcinus maenas. Brain Research 1025:85–97. Harrison, P.J.H., H.S. Cate, P. Steullet, and C.D. Derby. 2001a. Structural plasticity in the olfactory system of adult spiny lobsters: postembryonic development permits life-long growth, turnover and regeneration. Marine and Freshwater Research 52:1357–1365. Harrison, P.J.H., H.S. Cate, E.S. Swanson, and C.D. Derby. 2001b. Postembryonic proliferation in the spiny lobster antennular epithelium: rate of genesis of olfactory receptor neurons is dependent on molt stage. Journal of Neurobiology 47:51–66. Hartenstein, V. 2005. Development of insect sensilla. Pages 379–419 in L.I. Gilbert, K. Iatrou, and S.S. Gill, editors. Comprehensive molecular insect science, Vol. 1, Elsevier Pergamon, Oxford. Hartenstein, V., S. Spindler, W. Pereanu, and S. Fung. 2008. The development of the Drosophila larval brain. Pages 1–31 in G.M. Technau, editor. Brain development in Drosophila melanogaster. Landes Bioscience, Austin. Hartman, H.B., and R.L. Cooper. 1994. Regeneration and molting effects on a proprioceptor organ in the Dungeness crab, Cancer magister. Journal of Neurobiology 25:461–471. Hartnoll, R.G. 1985. Growth, sexual maturity and reproductive output. Pages 101–128 in A.M. Wenner, editor. Crustacean issues, Vol. 3, Factors in adult growth. A.A. Balkema, Rotterdam. Harzsch, S. 2003. Ontogeny of the ventral nerve cord in malacostracan crustaceans: a common plan for neuronal development in Crustacea, Hexapoda and other Arthropoda? Arthropod Structure and Development 32:17–37. Harzsch, S., and R.R. Dawirs. 1994. Neurogenesis in larval stages of the spider crab Hyas araneus (Decapoda, Brachyura): proliferation of neuroblasts in the ventral nerve cord. Roux’s Archives of Developmental Biology 204:93–100. Harzsch, S., and R.R. Dawirs. 1995/1996. Maturation of the compound eyes and eyestalk ganglia during larval development of the brachyuran crustaceans Hyas araneus L. (Decapoda, Majidae) and Carcinus maenas L. (Decapoda, Portunidae). Zoology 99:189–204. Harzsch, S., and R.R. Dawirs. 1996. Neurogenesis in the developing crab brain: postembryonic generation of neurons persists beyond metamorphosis. Journal of Neurobiology 29:384–398. Harzsch, S., and B.S. Hansson. 2008. Brain architechture in the terrestrial hermit crab Coenobita clypeatus (Anomura, Coenobitidae), a crustacean with good aerial sense of smell. BMC Neuroscience 9:58. Harzsch, S., J. Miller, J. Benton, and B.S. Beltz. 1999. From embryo to adult: persistent neurogenesis and apoptotic cell death shape the lobster deutocerebrum. Journal of Neuroscience 19:3472–3485. He, S., D. Nakada, and S.J. Morrison. 2009. Mechanisms of stem cell self-renewal. Annual Review of Cell and Developmental Biology 25:377–406. Henke, K. 1953. Über Zelldifferenzierung im Integument der Insekten und ihre Bedingungen. Journal of Embryology and Experimental Morphology 1:217–26. Kamiguchi, Y. 1972. On the sexual dimorphism in the antennule of the freshwater prawn, Palaemon paucidens. Zoological Magazine 81:223–226. Keil, T.A. 1997. Comparative morphogenesis of sensilla: a review. International Journal of Insect Morphology and Embryology 26:151–160. Keil, T.A., and R.A. Steinbrecht. 1984. Mechanosensitive and olfactory sensilla of insects. Pages 477–516 in R.C. King and H. Akai, editors. Insect ultrastructure, Vol. 2. Plenum, New York. Kriegstein, A., and A. Alvarez-Buylla. 2009. The glial nature of embryonic and adult neural stem cells. Annual Review of Neuroscience 32:149–184. Lavalli, K.L. 1990. Survival and growth of early-juvenile American lobsters Homarus americanus through their first season while fed diets of mesoplankton, microplankton, and frozen brine shrimp. Fishery Bulletin 89:61–68.

201

202

Manfred Schmidt Laverack, M.S. 1976. External proprioceptors. Pages 1–63 in P.J. Mill, editor. Structure and function of proprioceptors in the invertebrates. Chapman and Hall, London. Laverack, M.S. 1978. The organization and distribution of CAP organs in the lobster Homarus gammarus (L.). Marine Behaviour and Physiology 5:201–208. Laverack, M.S. 1988. Larval locomotion, sensors, growth and their implication for the nervous system. Symposium of the Zoological Society of London 59:103–122. Lazarini, F., and P.-M. Lledo. 2011. Is adult neurogenesis essential for olfaction? Trends in Neurosciences 34:20–30. Letourneau, J.G. 1976. Addition of sensory structures and associated neurons to the crayfish telson during development. Journal of Comparative Physiology A 110:13–23. Lindsey, B.W., and V. Tropepe. 2006. A comparative framework for understanding the biological principles of adult neurogenesis. Progress in Neurobiology 80:281–307. Linser, P.J., H.G. Trapido-Rosenthal, and E. Orona. 1997. Glutamine synthetase is a glial-specific marker in the olfactory regions of the lobster (Panulirus argus) nervous system. Glia 20:275–283. Macmillan, D.L., T. Stuart, and M. Thomas. 1998. Development of proprioceptive organ on the walking legs of the rock lobster Jasus edwardsii (Decapoda: Palinuridae) by ordered addition and loss of receptor elements. Journal of Crustacean Biology 18:1–9. Martin, J.W., and G.E. Davis. 2001. An updated classification of the recent Crustacea. Natural History Museum of Los Angeles County Contributions to Science 39:1–164. Mashaly, A., M. Winkler, I. Frambach, H. Gras, and F.-W. Schürmann. 2008. Sprouting interneurons in mushroom bodies of adult cricket brains. Journal of Comparative Neurology 508:153–174. Maxwell, K.E., T.R. Matthews, M.R.J. Sheehy, R.D. Bertelsen, and C.D. Derby. 2007. Neurolipofuscin is a measure of age in Panulirus argus, the Caribbean spiny lobster, in Florida. Biological Bulletin 213:55–66. McClure, K.D., and G. Schubiger. 2007. Transdetermination: Drosophila imaginal disc cells exhibit stem cell-like potency. International Journal of Biochemistry and Cell Biology 39:1105–1118. Mellon, DeF. Jr., H.R. Tuten, and J. Redick. 1989. Distribution of radioactive leucine following uptake by olfactory sensory neurons in normal and heteromorphic crayfish antennules. Journal of Comparative Neurology 280:645–662. Melzer, R.R., C. Michalke, and U. Smola. 2000. Walking on insect paths? Early ommatidial development in the compound eye of the ancestral crustacean, Triops cancriformis. Naturwissenschaften 87:308–311. Meyer, N.P., and E.C. Seaver. 2009. Neurogenesis in an annelid: characterization of brain neural precursors in the polychaete Capitella sp. I. Developmental Biology 335:237–252. Meyer-Rochow, V.B., D. Towers, and I. Ziedins. 1990. Growth patterns in the eye of Petrolisthes elongatus (Crustacea; Decapoda; Anomura). Experimental Biology 48:329–340. Morante, J., C. Desplan, and A. Celik. 2007. Generating patterned arrays of photoreceptors. Current Opinion in Genetics and Development 17:314–319. Morrison, S.J., and A.C. Spradling. 2008. Stem cells and niches: mechanisms that promote stem cell maintenance throughout life. Cell 132:598–611. Nilsson, D.E., and A. Kelber. 2007. A functional analysis of compound eye evolution. Arthropopod Structure and Development 36:373–385. Norenberg, M.D. 1979. The distribution of glutamine synthetase in the rat central nervous system. Journal of Histochemistry and Cytochemistry 27:756–762. Packard, A., and V. Albergoni. 1970. Relative growth, nucleic acid content and cell numbers of the brain in Octopus vulgaris (Lamarck). Journal of Experimental Biology 52:539–552. Parker, G.H. 1890. The histology and development of the eye in the lobster. Bulletin of the Museum of Comparative Zoölogy at Harvard College 20:1–60. Rakic, P. 2002. Adult neurogenesis in mammals: an identity crisis. Journal of Neuroscience 22:614–618. Ready, D.F., T.E. Hanson, and S. Benzer. 1976. Development of the Drosophila retina, a neurocrystalline lattice. Developmental Biology 53:217–240. Reddy, G.V., B. Gupta, K. Ray, and V. Rodrigues. 1997. Development of the Drosophila olfactory sense organs utilizes cell-cell interactions as well as lineage. Development 124:703–712.



Adult Neurogenesis in Crustaceans

Regier, J.C., J.W. Shultz, A. Zwick, A. Hussey, B. Ball, R. Wetzer, J.W. Martin, and C.W. Cunningham. 2010. Arthropod relationships revealed by phylogenomic analysis of nuclear protein-coding sequences. Nature 463:1079–1083. Sandeman, D., R. Sandeman, C. Derby, and M. Schmidt. 1992. Morphology of the brain of crayfish, crabs, and spiny lobsters: a common nomenclature for homologous structures. Biological Bulletin 183:304–326. Sandeman, D.C., J.L. Benton, and B.S. Beltz. 2009. An identified serotonergic neuron regulates adult neurogenesis in the crustacean brain. Developmental Neurobiology 69:530–545. Sandeman, D.C., F. Bazin, and B.S. Beltz. 2011. Adult neurogenesis: examples from the decapod crustaceans and comparisons with mammals. Arthropod Structure and Development 40:258–275. Sandeman, R.E., and D.C. Sandeman. 1996. Pre- and postembryonic development, growth and turnover of olfactory receptor neurons in crayfish antennules. Journal of Experimental Biology 199:2409–2418. Sandeman, R., and D. Sandeman. 2000. “Impoverished” and “enriched” living conditions influence the proliferation and survival of neurons in crayfish brain. Journal of Neurobiology 45:215–226. Sandeman, R.E., D. Clarke, D. Sandeman, and M. Manly. 1998. Growth-related and antennular amputation-induced changes in the olfactory centers of crayfish brain. Journal of Neuroscience 18:6195–6206. Schachtner, J., M. Schmidt, and U. Homberg. 2005. Organization and evolutionary trends of primary olfactory brain centers in Tetraconata (Crustacea+Hexapoda). Arthropod Structure and Development 34:257–299. Schmidt, M. 1997. Continuous neurogenesis in the olfactory brain of adult shore crabs, Carcinus maenas. Brain Research 762:131–143. Schmidt, M. 2001. Neuronal differentiation and long-term survival of newly generated cells in the olfactory midbrain of the adult spiny lobster, Panulirus argus. Journal of Neurobiology 48:181–203. Schmidt, M. 2002. Identification of neuroblasts that drive adult neurogenesis in the central olfactory pathway of decapod crustaceans. Society of Neuroscience Abstracts 28:7.4. Schmidt, M. 2007a. Identification of putative neuroblasts at the base of adult neurogenesis in the olfactory midbrain of the spiny lobster, Panulirus argus. Journal of Comparative Neurology 503:64–84. Schmidt, M. 2007b. The olfactory pathway of decapod crustaceans—an invertebrate model for life-long neurogenesis. Chemical Senses 32:365–384. Schmidt, M., and C.D. Derby. 2011. Cytoarchitecture and ultrastructure of neural stem cell niches and neurogenic complexes maintaining adult neurogenesis in the olfactory midbrain of spiny lobsters, Panulirus argus. Journal of Comparative Neurology 519:2283–2319. Schmidt, M., and W. Gnatzy. 1984. Are the funnel-canal organs the “campaniform sensilla” of the shore crab, Carcinus maenas (Decapoda, Crustacea)? II. Ultrastructure. Cell and Tissue Research 237:81–93. Schmidt, M., and S. Harzsch. 1999. Comparative analysis of neurogenesis in the central olfactory pathway of adult decapod crustaceans by in vivo BrdU labeling. Biological Bulletin 196:127–136. Schmidt, M., and DeF. Mellon, Jr. 2011. Neuronal processing of chemical information in crustaceans. Pages 123–147 in T. Breithaupt and M. Thiel, editors. Chemical communication in crustaceans. Springer, New York. Schmitz, B. 1992. Postembryonic development of the crayfish Procambarus clarkii and its tailfan mechanosensory system. Pages 69–90 in R.N. Singh, editor. Nervous systems, principles of design and function. Wiley Eastern Limited, New Dehli. Scholtz, G. 1992. Cell lineage studies in the crayfish Cherax destructor (Crustacea, Decapoda): germ band formation, segmentation, and early neurogenesis. Roux’s Archives of Developmental Biology 202:36–48. Shabani, S., M. Kamio, and C.D. Derby. 2008. Spiny lobsters detect conspecific blood-borne alarm cues exclusively through olfactory sensilla. Journal of Experimental Biology 211:2600–2608.

203

204

Manfred Schmidt Sheehy, M.R.J., R.C.A. Bannister, J.F. Wickins, and P.M.J. Shelton. 1999. New perspectives on the growth and longevity of the European lobster (Homarus gammarus). Canadian Journal of Fisheries and Aquatic Sciences 56:1904–1915. Shen, Q., Y. Wang, E. Kokovay, G. Lin, S.-M. Chuang, S.K. Goderie, B. Roysam, and S. Temple. 2008. Adult SVZ stem cells lie in a vascular niche: a quantitative analysis of niche cell-cell interactions. Cell Stem Cell 3:289–300. Siegrist, S.E., N.S. Haque, C.-H. Chen, B.A. Hay, and I.K. Hariharan. 2010. Inactivation of both foxo and reaper promotes long-term adult neurogenesis in Drosophila. Current Biology 20:643–648. Song, C.-K., L.M. Johnstone, M. Schmidt, C.D. Derby, and D.H. Edwards. 2007. Social domination increases neuronal survival in the brain of juvenile crayfish Procambarus clarkii. Journal of Experimental Biology 210:1311–1325. Song, C.-K., L.M. Johnstone, D.H. Edwards, C.D. Derby, and M. Schmidt. 2009. Cellular basis of neurogenesis in the brain of crayfish, Procambarus clarkii: neurogenic complex in the olfactory midbrain from hatchlings to adults. Arthropod Structure and Development 38:339–360. Steullet, P., H.S. Cate, and C.D. Derby. 2000. A spatiotemporal wave of turnover and functional maturation of olfactory receptor neurons in the spiny lobster Panulirus argus. Journal of Neuroscience 20:3282–3294. Stollewerk, A. 2008. Evolution of neurogenesis in arthropods. Pages 359–380 in A. Minelli and G. Fusco, editors. Key themes in evolutionary developmental biology. Cambridge University Press, Cambridge. Strausfeld, N.J. 2009. Brain organization and the origin of insects: an assessment. Proceedings of the of the Royal Society B: Biological Sciences 276:1929–1937. Strausfeld, N.J., I. Sinakevitch, S.M. Brown, and S.M. Farris. 2009. Ground plan of the insect mushroom body: functional and evolutionary implications. Journal of Comparative Neurology 513:265–291. Stuart, T., and D. Macmillan. 1997. Development of spines and sensory setae on the tailfan of the southern rock lobster (Jasus edwardsii Hutton, 1875) (Crustacea: Decapoda). Australian Journal of Zoology 45:307–315. Sullivan, J.M., and B.S. Beltz. 2005a. Adult neurogenesis in the central olfactory pathway in the absence of receptor neuron turnover in Libinia emarginata. European Journal of Neuroscience 22:2397–2405. Sullivan, J.M., and B.S. Beltz. 2005b. Newborn cells in the adult crayfish brain differentiate into distinct neuronal types. Journal of Neurobiology 65:157–170. Sullivan, J.M., J.L. Benton, D.C. Sandeman, and B.S. Beltz. 2007a. Adult neurogenesis: a common strategy across diverse species. Journal of Comparative Neurology 500:574–584. Sullivan, J.M., D.C. Sandeman, J.L. Benton, and B.S. Beltz. 2007b. Adult neurogenesis and cell cycle regulation in the crustacean olfactory pathway: from glial precursors to differentiated neurons. Journal of Molecular Histology 38:527–542. Tadesse, T., M. Schmidt, W.W. Walthall, P.C. Tai, and C.D. Derby. 2011. Distribution and function of splash, an achaete-scute homolog in the adult olfactory organ of the Caribbean spiny lobster Panulirus argus. Developmental Neurobiology 71:316–335. Taupin, P. 2007. BrdU immunohistochemistry for studying adult neurogenesis: paradigms, pitfalls, limitations, and validation. Brain Research Reviews 53:198–214. Tavazoie, M., L. Van der Veken, V. Silva-Vargas, M. Louissaint, L. Colonna, B. Zaidi, J.M. Garcia-Verdugo, and F. Doetsch. 2008. A specialized vascular niche for adult neural stem cells. Cell Stem Cell 3:279–288. Ungerer, P., and G. Scholtz. 2008. Filling the gap between identified neuroblasts and neurons in crustaceans adds new support for Tetraconata. Proceedings of the Royal Society B: Biological Sciences 275:369–376. Ungerer, P., B.J. Eriksson, and A. Stollewerk. 2011. Neurogenesis in the water flea Daphnia magna (Crustacea, Branchiopoda) suggests different mechanisms of neuroblast formation in insects and crustaceans. Developmental Biology 357:42–52. Williamson, D.I. 1982. Larval morphology and diversity. Pages 43–110 in L.G. Abele, editor. The biology of Crustacea, Vol. 2, Embryology, morphology, and genetics. Academic Press, New York. Wolff, T. 1978. Maximum size of lobsters (Homarus) (Decapoda, Nephropidae). Crustaceana 34:1–14.



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Zhang, Y., S. Allodi, D.C. Sandeman, and B.S. Beltz. 2009. Adult neurogenesis in the crayfish brain: proliferation, migration, and possible origin of precursor cells. Developmental Neurobiology 69:415–436. Zhang, Y., J.L. Benton, and B.S. Beltz. 2011. 5-HT receptors mediate lineage-dependent effects of serotonin on adult neurogenesis in Procambarus clarkii. Neural Development 6:2. Zhao, X., V. Coptis, and S.M. Farris. 2008. Metamorphosis and adult development of the mushroom bodies of the red flour beetle, Tribolium castaneum. Developmental Neurobiology 68:1487–1502.

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8 VISUAL SYSTEMS OF CRUSTACEANS

Raymon Glantz

Abstract This chapter examines the properties of the dioptic apparatus in compound and nauplius eyes, photoreceptor physiology, and the structure-function relationships of the neural pathways in the optic lobe and the brain. The discussion of the dioptic apparatus focuses on the structure of the ommatidium and the structural and geometric variations involved in the control of sensitivity and spatial resolution. The review of photoreceptors encompasses the formation of the rhabdom, structural attributes of photoreceptors related to photon capture and polarization vision, the photopigments and spectral sensitivity, phototransduction, and the dynamic properties of the light response. Four issues are central to the discussion of the neural mechanism in the optic lobe: (1) The role of lamina monopolar neurons in the establishment of the retinotopic columnar projection system; (2) The evidence that lamina monopolar cells provide separate channels for contrast and polarization vision (and possibly color vision); (3) The structure and physiology of the large-field identified neurons in the medulla externa and the lobula and related results suggesting that contrast vision and motion detection are partitioned between the medulla and lobula respectively; and (4) The functions of these neurons in optomotor reflexes and escape and defense behavior. The review concludes with a brief description of the visual interneurons and optomotor neurons in the protocerebrum of the brain and the evidence for extraretinal photoreceptors.

INTRODUCTION The study of visual systems is guided by two related questions. What is perceived, and how is it perceived? The “what” question is expressed in the relationship between an organism and its 206



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environment (see ­chapters 9 and 18 in this volume) and the interactions between conspecifics (see ­chapters 17–19 in this volume). This chapter addresses the how question, that is, the structure-function relationships that select, amplify, and filter the stimuli between the visual environment and behavior. This review begins with the dioptric apparatus and photoreceptor cells, examines signal transformations in the optic lobe and brain, and ends with the caudal photoreceptor and extraretinal photoreceptors. The discussion of peripheral mechanisms focuses on the control of visual sensitivity and the basis of polarization and spectral sensitivity. Studies on optic lobe focus on information processing related to behaviorally relevant stimuli. Comparative data are examined on the optics and retinal structures for a wide variety of crustaceans. Physiological investigations are focused entirely on decapods and barnacles (Balanus).

OMMATIDIA The majority of crustacean groups have compound eyes either as adults or in larval stages. Eye structures in diurnal and terrestrial forms tend to maximize resolution within the limits of a discretized dioptric system. Conversely, in deep-sea forms, the eyes are often specialized to capture photons at the expense of resolution. All compound eyes are composed of ommatidia with as few as four in some isopods to over 10,000 in adult clawed lobsters (Homarus). In contrast to flies, the ommatidium is the optical unit of all crustacean compound eyes, that is, all the photoreceptor cells in an ommatidium share the same optic axis. The dioptric apparatus fulfills two essential requirements of vision. First, the corneal facet collects light over a relatively large area (when compared to the photoreceptor cross section) and focuses it to a smaller region, the distal tip of the rhabdom (a local aggregate of photoreceptors). Second, light from different directions is captured by different ommatidia, which is the basis for spatial vision (as shown in Fig. 8.1). Earlier work on ommatidial structure is reviewed in Land (1981, 1984), Shaw and Stowe (1982), and Land and Nilsson (2006). The main components of the ommatidium are a corneal facet, a crystalline cone, and a rhabdom, all aligned on the optic axis. The crystalline cone and the rhabdom are surrounded by accessory pigment cells. Shielding pigments may also occupy the cytoplasm of retinular cells (photoreceptors). In crustaceans with superposition eyes (see below), the pigments are redistributed during light and dark adaptation and control visual sensitivity by absorbing off-axis illumination. The base of each ommatidium may contain a tapetum, which reflects light back through the retinula, increasing the photon catch. Corneal facets are typically hexagonal but are square in three groups of macruran decapods and galatheid squat lobsters. The facet lens often lacks sufficient power to focus light on the rhabdom. Refractive power then resides in the crystalline cones, which form lens cylinders (Shaw and Stowe 1982, Land 1984). The lens cylinder is formed by a radially symmetric and graded index of refraction; that is maximal at the center and tapering off toward the margin. The length of the cylinder determines the location of the image plane, which varies with the type of compound eye. The cone may be connected to the rhabdom through a crystalline fiber. The main rhabdom is composed of 5 to 7 rhabdomeres (typically 7 in decapods and stomatopods) composed of retinular cell microvilli. An eighth retinular cell in decapods, stomatopods, and mysids often forms a separate small rhabdom distal to the main rhabdom. Homologs of the eighth receptor cell are also present in branchiopods, branchiurans, and amphipods. The principal function of the rhabdom is photon absorption and visual transduction but it is also a light guide. The cross-sectional diameter of the rhabdom affects photon capture and thus sensitivity. In a number of crabs (Ocypode, Uca, Leptograpsus), the distal rhabdom diameter can be as small as 2–3 µm (Shaw and Stowe 1982, Waterman 1984) compared with about 20 µm in the nocturnal

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Raymon Glantz A

Image

B

Image

cf

cc cz rh bm ax

Fig. 8.1. Schematic representation of a generic (A) apposition eye and (B) reflecting superposition eye. The broken lines indicate light rays and the heavy lines between the ommatidia indicate accessory shielding pigments. An important feature of the apposition eye (A)  is that off-axis light rays are absorbed by the shielding pigments, but in the superposition eye (B) the off-axis light rays are reflected from the walls of the crystalline cones and enter neighboring ommatidia through the clear zones. The single axon at the base of each ommatidium is representative of a bundle of eight axons. Abbreviations: ax, receptor axons; bm, basement membrane; cc, crystalline cones; cf, corneal facets; cz, clear zone; rh, rhabdom.

crayfish Procambarus. The axons of the retinular cells exit the ommatidium via the basement membrane en route to the lamina ganglionaris (first optic neuropil).

RHABDOM AND RETINULAR CELL STRUCTURE With only a few exceptions, crustacean ommatidia have closed rhabdoms (Waterman 1981); that is, the space between adjacent retinular cells is occluded by the microvilli. In decapods, stomatopods, mysids, and euphausids, the rhabdom is fused and banded due to the interdigitation of orthogonal microvilli from several retinular cells. Variants include Panulirus and the mysid Mysidium gracile, in which only a portion of the rhabdom is banded. In many branchiopods (e.g., Daphnia and Artemia) and amphipods, the rhabdom is closed but unbanded. An interesting exception is the cyclopean eye of the water-flea Polyphemus. The 130 ommatidia exhibit four different types of partially banded rhabdoms in distinct retinal zones (Odselius and Nilsson 1983). In decapods, the long axes of retinular cell microvilli form parallel arrays perpendicular to the optic axis. The orientation of microvilli is essential to arthropod polarization sensitivity and it is considered further in that context. The lengths of microvilli vary from 1 to 2 μ m in a variety of crabs, for example, Uca, Leptograpsus, Grapsus, and Callinectes (Shaw and Stowe 1982), to about 8 μ m in crayfish. Microvilli diameters are on the order of 60 to 90 nm. The core of the microvillus contains actin filaments (Matsushita and Arikawa 1997), and the plasma membrane is packed with rhodopsin particles of about 100-nm diameter (Eguchi et al. 1989).



Visual Systems of Crustaceans

COMPOUND EYES Compound eyes are present in most crustacean subclasses. The exceptions are copepods, the majority of ostracods, cephalocarids, and adult barnacles. In stalk-eyed crustaceans, the cornea is roughly hemispherical with a panoramic visual field. Apposition and superposition eyes differ in the organization of the dioptric pathway, and this difference has consequences for sensitivity and spatial resolution. Apposition eyes are characteristic of terrestrial and diurnal organisms, and superposition eyes are more common among nocturnal and deep-sea species.

APPOSITION EYES Apposition eyes (as in Fig. 8.1A) are characteristic of a large majority of crustaceans and found in decapods, branchiopods, isopods, amphipods, stomatopods, brachyurans, and most anomurans. In apposition eyes, the ommatidia are optically isolated from one another by accessory pigment cells and/or screening pigment within the retinular cells. Each corneal facet subtends a small solid angle of visual space, and the dioptric apparatus forms an inverted image of this space on the distal surface of the rhabdom. Although the local image is inverted, the global image is erect. Details of the local image do not contribute to form vision. All of the photoreceptor cells in a given rhabdom receive the same stimulus. The rhabdom lies just proximal to the crystalline cone. The refractive power of the dioptric apparatus is critical. In a typical diurnal crab, the shore crab, the light captured by a 20- μ m diameter corneal facet must be focused on a rhabdom tip of about 2 to 5 μ m diameter. The length of the crystalline cone in apposition eye is equal to the focal length, which ensures that the image plane lies at its proximal tip. Apposition eyes may reside on mobile eyestalks as in the true crabs or sessile structures as in some amphipods. They also vary widely in the size of the visual field, the number of ommatidia, and the visual functions they support. Thus an amphipod (e.g., sandhoppers) with about 300 ommatidia in each eye can use vision for solar and lunar orientation. The cyclopean eye of Daphnia is small, but the eye’s mobility extends the visual space to over 100°.

ACUTE ZONES Acute zones are common in apposition eyes and provide a mechanism to enhance spatial resolution (reviewed in Land 1981). Compound eyes have limited resolution compared with vertebrate eyes: about 0.5 to 1.0 cycle/deg compared to 40 cycles/deg in human eyes. The limited resolution results from a combination of the large panoramic field of view, small eye size, and the division of visual space by discrete corneal facets. Thus a compound eye with 100 facets in each horizontal row and a visual field of 180°, will have an average interommatidial angle (Δφ) of 1.8° and a spatial resolution (1/2 Δφ) of about 0.28 cycles/deg. The requirements of spatial resolution place a premium on the curvature of the cornea. Littoral crabs use vision to forage, defend their burrows, mate, and avoid predation (Zeil and Al-Mutairi 1996). Acute zones are corneal areas with enlarged facets and f lattened contour so as to reduce Δφ. In the ghost crab (Ocypode) and fiddler crabs (Uca) (Zeil and Al-Mutairi 1996, Smolka and Hemmi 2009), the eyestalks are held vertically erect. A horizontal band of corneal facets along the eye equator is enlarged relative to the vertically adjacent neighbors. In these crabs, spatial resolution in this zone is about 1 to 2 cycles/deg compared to 0.2 to 0.3 cycles/deg in other areas of the

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Raymon Glantz eye. The corneal facets in the acute zone are about twice the diameter of facets of the dorsal cornea. Within the acute zone, vertical resolution may be twice that parallel to the horizon. In several marine crustaceans acute zones are implemented in divided eyes, that is, eyes with two separate spherical segments. Thus in the apposition eyes of the hyperiid amphipod, Hyperia, Δφ = 0.8º in the dorsal eye and 10.5º in the ventral eye. The rationale for this asymmetry is that only the dorsal visual field has sufficient illumination for vision. Divided eyes with similar features are also found in the superposition eyes of mysids and euphausids (Land 1989, Land and Nilsson 2006). In the mysid shrimp Dioptromysis, the main lobe of the eye has superposition ommatidia and 16-μm diameter facets. The back of the eye consists of a single ommatidium with a 44-μm facet illuminating 120 rhabdoms. In deep-water euphausids, the dorsal facets are larger and the visual field is narrower than the ventral ommatidia. In transparent organisms including diverse zooplankton, hyperiid amphipods and the larval stages of euphausids, stomatopods, and decapods, the accessory screening pigments are absent. To maintain optical isolation between ommatidia, the hyperiid amphipod Phronima and decapod larvae combine an elongated crystalline cone-lens cylinder and a crystalline fiber light guide. In larval euphausids, the proximal end of the crystalline cone forms a second lens, which restricts light to the ommatidial axis.

STOMATOPOD EYES Stomatopods are diurnal predators, and their eyes exemplify the evolution and specialization of apposition eyes. Their eyes have an unusual global geometry and receptor cell organization (reviewed in Shaw and Stowe 1982, Cronin 2006, ­chapter 9 in this volume). The eyes consist of dorsal and ventral spherical segments (“hemispheres”) separated by a midband consisting of six rows of ommatidia. The two hemispheres are similar to those of decapods (similar dioptrics and rhabdom structure) but with exceptional curvature. In the region where the hemispheres approach the midband, the corneas curve inward. Pseudopupil measurements near the equator of the eye reveal three dark zones: one in each of the hemispheres and one in the midband. Thus facets in the two hemispheres share optical axes with midband facets. The ommatidia of the hemispheres include hexagonal facets, a crystalline cone, and eight retinular cells. Seven retinular cells form a fused and banded rhabdom, and the eighth is a violet receptor cell forming a separate, distal rhabdom. This pattern is similar to the organization of most decapod ommatidia. The midband rows have enlarged facets (135 μm in diameter), small Δφ (0.5º), and a tiered rhabdom. The retinular cells in the midband are specialized for color and polarization vision. The visual capabilities of these remarkable eyes are extended by frequent eyestalk movements, which are used to continuously track objects. The optical requirements for high sensitivity are challenging for apposition eyes and produce unusual outcomes in deep-sea organisms. The general solution is to sacrifice acuity with large corneal facets, thick and tapering crystalline cones, thick rhabdoms, and/or a reflecting tapetum. Various combinations of these strategies are employed in the sessile eye of the amphipod Phronima (Land 1981), the deep-sea isopod Cirolana borealis, and the deep-sea half-crab Paralomis multispina (Eguchi et al. 1997).

SUPERPOSITION EYES Superposition eyes (as in Fig. 8.1B) are optimized for sensitivity. They form a single erect image of visual space on the retina. The light that forms each point of the focused image enters the eye



Visual Systems of Crustaceans

through numerous corneal facets. A good image requires a homogeneous array of facets and near identical Δφ across the cornea. Superposition eyes are found in macruran decapods, mysids, euphausids, and galatheid anomurans. Some forage at night (e.g., crayfish) or spend their days in the ocean depths (euphausids and some mysids). In a superposition eye in the dark-adapted state, the effective pupil area is up to 10 ommatidia wide (Shaw and Stowe 1982). This implies a 100-fold increase in the rate of photon capture. Furthermore, high sensitivity in superposition eyes is supported by a thick rhabdom (20 μm in Procambarus clarkii and 35 μm in Cherax destructor). Since the receptor cell’s acceptance angle Δp is proportional to the rhabdom diameter and sensitivity varies as Δp2, the rhabdom diameter makes a substantial contribution to sensitivity. High sensitivity also requires a large diameter lens and a short focal length. These result in a rapidly converging light cone at the proximal end of the dioptric apparatus. Thus one function of superposition optics is to collimate the light. A defining feature of these eyes is a clear zone between the proximal tip of the crystalline cone and the rhabdom. The crystalline cones of superposition eyes are longer than in apposition eyes, typically twice their focal lengths. The image formed by the dioptric apparatus of superposition eyes may depend on refraction, reflection, or a combination of these (Land 1989). Reflecting superposition eyes are unique to macruran decapods and galatheid squat lobsters. The corneal facets are square, and the crystalline cone has a square cross section (i.e., a pyramid) lined with mirrored walls. The mirrors produce a corner reflection for paraxial light rays, and the light ray emanating from the crystalline cone lies in the same plane as the incident ray but with a reversed direction. Strictly axial rays are undeviated. The doubly reflected rays cross the clear zone at an angle to the optic axis and enter neighboring ommatidia. The ray path requires that the accessory screening pigments have vacated the areas adjacent to the clear zone. During light adaptation, proximal screening pigments move distally and the distal pigments move proximally surrounding the clear zone. The acceptance angles of retinular cells can narrow by three- to six-fold in crayfish (Shaw and Stowe 1982). Refracting superposition eyes are characteristic of euphausids (krill) and mysids. The optics are based on the refractive power of the crystalline cone lens cylinder. The length of the cone is twice its focal length, and the emerging rays are effectively collimated and afocal (Land 1984, Land and Nilsson 2002). The effect of the lens cylinder is to focus the light at a focal plane in the middle (lengthwise) of the cone and to collimate it in the proximal half of the lens while bending the rays to cross the optic axis. The most recent addition to this family, the parabolic superposition eye, combines reflection and refraction (Land and Nilsson 2006) and is present in several crabs and hermit crabs. The essential feature is a crystalline cone with a mirrored and inwardly curving parabolic profile. The cone produces collimated light.

SIMPLE EYES Most crustaceans go through a larval stage identified by a characteristic unpaired nauplius eye on the midline of the head. But for the copepods, at least some members of all other subclasses develop compound eyes (Nilsson 2009). Compound eyes may replace the nauplius eye or develop alongside it. In addition to the copepods, nauplius eyes persist in the adults of most branchiopods, ostracods, and cirripedes. Compound eyes and nauplius eyes arise from different embryonic structures. The planktonic forms exhibit light-sensitive vertical migrations, and several species, for example Pontella karachiensis, are sensitive to polarized light (Manor et al. 2009). In the majority of copepods, the nauplius eye is a tripartite structure consisting of three small adjacent eyecups, each with a small number of rhabdomeric receptors. There are no lenses, and the only optical refinement may be a reflecting tapetum lining the surface of the eyecup

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Raymon Glantz beneath the photoreceptors. The eyes have little or no spatial resolution and appear to support phototaxis. However, there are a good number of exceptions described by Land (1981, 1984) and Land and Nilsson (2006) among the pontellid copepods and ostracods. Ostracods may have compound eyes (e.g., Macrocypridina castanea) or simple eyes (e.g., in Gigantocypris) (Land 1984) or both as in the Myodocopida, Skogsbergia lerneri (Oakley and Huber 2004), and they inhabit nearly every aquatic niche. The deep-sea ostracod Gigantocypris has two nauplius eyes, 3.0 mm across. The back of each eye is covered with a reflecting tapetum. The tapetum functions as a light collector and concentrates light on a receptor area consisting of about 1,000 photoreceptor cells. Pontellid copepods have adapted several strategies based on the separation of the three nauplius eyecups with doublet or even triplet lenses (Land 1984). In Copilia, a pair of eyes are situated laterally in the head, and the surface of each eye contains a biconvex lens with a long focal length. There is a smaller but more powerful lens in the unpaired ventral eye. The latter focuses light on four to seven receptor cells. The eyes have only a small window on visual space (about 3º) but extend the field by constant scanning motions. In Labidocera, the two dorsal eyes are joined and the retinular cells (five from each eye) form a linear array. The receptor array subtends 30–40º laterally, but only a few degrees along the anterior-posterior axis. To compensate, the eye scans a 40º field along the anterior-posterior axis. Furthermore, eye movements will track overhead illumination (Land 1988) in a manner reminiscent of the dorsal light reflex in decapods. In P. karachiensis, the retinular cells have parallel microvilli, and the microvilli of at least some of the adjacent receptors are orthogonal (Manor et al. 2009). A new class of simple eye has been discovered in bresiliid shrimps Rimicaris exoculata and Chorocaris chacei (Lakin et al. 1997), which inhabit hydrothermal vents along the mid-Atlantic ridge. At these depths, there is no downwelling light and compound eyes are absent. A pair of simple eyes sits atop the cephalic carapace caudal to the rostrum. The surface of the eye is a 2- to 3-mm oval of transparent cuticle. The retina consists of a high density of rhabdoms, each composed of five retinular cells with parallel microvilli. Rhabdom density in C. chacei is about six times that in the shrimp Palaemonetes. It is suggested that the eyes detect infrared light from the hydrothermal vents. Barnacles also have simple eyes. The medial ocellus contains four to seven photoreceptor cells, and the lateral ocelli have three receptor cells. Each receptor cell has a distal dendrite with microvilli. The medial receptor cells have large-diameter axons that project directly to the supraesophageal ganglion. The receptor potential is transmitted decrementally via the receptor axons to the synaptic terminals (Shaw and Stowe 1982).

SCREENING PIGMENTS AND PHOTORECEPTOR MEMBRANE CYCLING Sensitivity of retinular cells is influenced by the circadian turnover of microvilli (Eguchi 1999), the migration of shielding pigments in retinular and accessory cells, and modulation of the transduction pathway. These mechanisms are expressed to different degrees among various crustaceans. Migration of accessory pigment controls the amount of light that reaches the rhabdom (Rodríguez-Sosa and Aréchiga 1982, Meyer-Rochow 2001). The distal pigments are contained in accessory cells housed along the partition between the ommatidia, and the proximal pigments are contained within the retinular cells. Acting together, the pigments form a longitudinal pupil regulating the transmission of off-axis light rays (Bryceson and McIntyre 1983, King and Cronin 1994). The distal screening pigment determines the retinular cell’s acceptance angle. In superposition eyes, the acceptance angle is diminished by 65–75% during light adaptation (Shaw and Stowe 1982). This migration narrows the rhabdom’s aperture from 35 μm in the dark-adapted eye



Visual Systems of Crustaceans

to 5 μm in a light-adapted state. In this condition, the superposition eye is functionally similar to an apposition eye. In addition to controlling the quantum catch, screening pigments alter the spectrum of the transmitted radiation (Goldsmith 1978), producing a red-shift of 30–35 nm. In the apposition eye of Ligia, the pigments of the accessory cells do not migrate (Hariyama et al. 2001) and the acceptance angle of retinular cells (about 2º) is invariant. Sensitivity is controlled by dispersal of the retinular cell’s screening pigment. Membrane recycling occurs in a circadian cycle. The distal tips of rhabdoms are removed by phagocytosis, and new rhodopsin is generated proximally in the soma (reviewed in Eguchi 1999). A specialized transport system conveys a rhodopsin-membrane complex to the base of the rhabdomere, where new microvilli are generated. The membrane recycling also produces a circadian rhythm in photoreceptor sensitivity (reviewed in Aréchiga and Rodríguez-Sosa 2002).

PHOTOPIGMENTS The absorption of light by a photopigment is the sine qua non of vision. Crustacean visual pigments (reviewed by Cronin and Hariyama 2002, Cronin 2006) share many features with the rhodopsins of the vertebrate retina. The rhodopsins are based on opsin proteins bound to a chromophore, most commonly retinal (A1 rhodopsin) or less often 3,4-dehydroretinal (A2 rhodopsin). Both the chromophore and the opsin influence the absorption spectra. In general, the λ max of A2 rhodopsins are red-shifted relative to A1 rhodopsins. The chromophore can vary seasonally. A distinctive feature of arthropod rhodopsins concerns the disposition of the chromophore following excitation. Photon absorption isomerizes 11-cis-retinal to the all-trans configuration that in turn triggers the thermal conversion of rhodopsin to metarhodopsin. In arthropods, metarhodopsin is relatively thermostable and ambient illumination can regenerate the 11-cis isomer and native rhodopsin. In vertebrates, the retinal isomer is rapidly separated from the opsin. Because the 11-cis and all-trans isomers have different absorption spectra, the initial isomerization and the regeneration have different λ max values. In addition to the binding pocket for the chromophore, opsin proteins have seven hydrophobic transmembrane segments and an intracellular binding site for the G-protein (Sakamoto et al. 1996, Porter et al. 2007). The analysis of the absorption spectra of crustacean rhodopsins and the spectral sensitivity profiles of retinular cells are motivated by two major issues: adaptation to the local irradiance spectrum (see ­chapters 9 and 10 in this volume) and color vision. The study of color vision has largely focused on two questions: (1) Does an organism have retinular cells containing photopigments with different absorption spectra? and (2) Can an organism distinguish objects based exclusively upon the spectrum of reflected light? Absorption spectra are typically measured with microspectrophotometry (MSP) or the electroretinogram (ERG). While MSP can resolve the spectrum of single cells, the ERG may fail to capture the spectrum of pigments present in only a minority of receptor cells, for example, R8 in many crabs (Jordão et al. 2007). Furthermore, the ERG cannot indicate whether two spectral peaks are derived from distinct photoreceptor cells. A third method is to measure the spectral sensitivity profile of single retinular cells using intracellular recording. There is solid evidence that a number of crustaceans have two or more photopigments located in different retinular cells. Crayfish are dichromats with a green (λ max = 530 nm) absorbing pigment in the main rhabdom and a violet sensitive pigment (λ max = 440 nm) in the distal rhabdomere of R8 (Cummins and Goldsmith 1981). The blue-green absorption dichotomy found in Procambarus appears to occur in a number of decapods including other crayfish species, several groups of crabs (Martin and Mote 1982, Knight and Leggett 1985, Horch et al. 2002), spiny lobsters (Cummins et al. 1984), fresh-water shrimp Palaemonetes (Goldsmith and Fernandez 1968), and deep-sea shrimp (Frank and Case

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Raymon Glantz 1988). In a number of crustaceans, the short-wavelength sensitivity peaks in the ultraviolet. Neurophysiological data in crayfish (reviewed in Shaw and Stowe 1982) suggest that the two chromatic channels converge on the sustaining fibers (SusF) of the medulla externa (second optic neuropil). There is no evidence of spectrally selective inhibition (opponency). The optic lobe and brain of Paragrapsus (Knight and Leggett 1985) contain ultraviolet- and green-­sensitive neurons as in retinular cells, but there are also yellow- and blue-sensitive interneurons that may reflect the filtering effects of screening pigments. There is no indication of opponency. Behavioral studies suggest that several groups of presumptive dichromatic decapods may possess true color vision (Waterman 1961, Detto et al. 2006, Detto 2007), including fiddler crabs, the sand shrimp Crangon crangon, Carcinus, and the amphipod Caprella liparotensis. Stomatopods provide the strongest case for color vision with 10 or more photopigments distributed in different photoreceptor cells (see ­chapter 9 in this volume) and the capacity to distinguish objects based on the reflected spectrum (Marshall et al. 1996). A number of crustaceans have multiple photopigments, including the isopod Ligia exotica (Hariyama et  al. 1993), ostracods (Oakley and Huber 2004), and Daphnia. In Daphnia, each ommatidium contains eight retinular cells, divided into four subsets distinguished by their spectral sensitivity (Smith and Macagno 1990) with λ max values of 348, 434, 525, and 608 nm. Thus Daphnia may be unique among crustaceans in that individual rhabdoms contain retinular cells with different photopigments. Molecular studies in other branchiopods (Kashiyama et al. 2009) have revealed up to five opsin genes in a single species. Daphnia uses its spectral sensitivity to execute a set of wavelength-specific phototactic responses, color dances, that discriminate long versus short and are associated with feeding on phytoplankton.

POLARIZATION SENSITIVITY Interest in polarization sensitivity (PS) was initially motivated by von Frisch’s studies of navigation in the honey bee. Polarization sensitivity is a hallmark of microvillar photoreceptor cells. If the microvilli of a receptor cell are parallel to one another, the receptor will be maximally sensitive to the electric vector (e-vector) of light that oscillates in a parallel plane. The magnitude of the polarization sensitivity should ref lect the degree to which the light-absorbing (i.e., dichroic) axes of the rhodopsin molecules are aligned along the axis of the microvilli. Polarization sensitivity has been described in a wide variety of insects, crustaceans, and cephalopods (reviewed in Waterman 1981, Shaw and Stowe 1982). Among crustaceans, PS is present for linearly polarized light in decapods, stomatopods, mysids, and in a several branchiopods and copepods (Manor et al. 2009). In Daphnia pulex, behavioral responses to polarized light exhibit a complex wavelength dependence (Novales Flamarique and Browman 2000). Polarized light is described by three parameters: intensity, e-vector orientation (θ), and the degree of polarization. Photoreceptor cells are characterized by the magnitude of the PS and the e-vector orientation of maximum sensitivity (θ max). In decapods and stomatopods, PS magnitudes typically vary between 3.0 and 9.0. The high PS implies that the dichroic axis of rhodopsin is at least partially aligned with the long axis of the microvillus. A random orientation would produce a dichroic ratio about 2.0 (Goldsmith 1975, reviewed in Waterman 1981, Shaw and Stowe 1982). In decapods, the main rhabdom is composed of seven retinular cells characterized by two geometries with orthogonal microvilli (Waterman 1981) and thus θ max values come in pairs separated by 90º. Furthermore, in the dorsomedial section of the crayfish eye, a substantial number of ommatidia are rotated by about 45º, and these ommatidia exhibit θ max pairs of 0 and 90º or 45 and 135º (Glantz 2007).



Visual Systems of Crustaceans

The threshold for PS (the minimum degree detectable in behavioral responses, see below) is 10 to 20% degree of polarization. Stomatopod eyes are exceptional in many respects, not least of which is in PS (Kleinlogel and Marshall 2006). In midband rows 5 and 6, the main rhabdom is specialized for PS. Furthermore there is PS in the adjacent hemispheric photoreceptor cells but with a different λ max and with θ max rotated by 45º relative to the midband. Finally R8 in midband rows 5 and 6 also exhibits PS in the ultraviolet (Cronin et al. 1994, Kleinlogel and Marshall 2009). The midband R8 cells exhibit a 90º difference in θ max between rows 5 and 6. The data suggest that some stomatopods may be able to simultaneously analyze the light impinging on the midband and adjacent hemispheres into four e-vector channels at three wavelengths. Furthermore, species of Odontodactylus are uniquely sensitive to circularly polarized light (Chiou et al. 2008). In midband rows 5 and 6, the optical properties of the R8 photoreceptor cell convert circular polarization to linear polarization. The R1-R7 receptor cells respond selectively to circularly polarized light but not to linearly polarized light.

THE PHOTORECEPTOR CELL’S LIGHT RESPONSE AND ADAPTATION All of the above issues influence the retinular cell’s response to light, and the photoreceptor membrane adds additional operators to visual function. These include contributions to light and dark adaptation, temporal integration, and dynamic behavior in response to time varying stimuli. Arthropod retinular cells respond to increments of illumination with depolarization (as in Fig. 8.2A). Response magnitude and timing depend on both intensity and the state of adaptation. Low intensity steps of light (< 10 times threshold) elicit slowly rising responses that reach a plateau (typically < 10 mV) in several hundred msec and maintain a constant depolarization for the stimulus duration. In this intensity range, the response magnitude is approximately linear with light intensity and there is little evidence of adaptation. This intensity domain supports the range of spatial contrast in natural scenes. At higher intensities, the step response is faster (time to peak may be 500 nm in the shallowest-living species down to nearly 470 nm in the deepest-living ones. The other retinal regions display no obvious trend of visual pigment λ max with habitat. Nevertheless, stomatopod retinas are adapted to their photic environment by the filters that tune photoreceptor spectral sensitivity. The traces in Fig. 9.5A suggest how much the filter absorbance spectra can vary among species, and Fig. 9.5C displays sensitivity maxima Fig. 9.5. (Continued) photoreceptors, tuning spectral sensitivity spectra of these receptors. (B) Visual pigments in 17 species of mantis shrimps. Species with fewer than four classes of filters are plotted with crosses through the symbol. Each point marks the λ max of the visual pigment in the receptor class at the retinal location indicated on the x-axis in one species. Ommatidial rows in the equatorial band are indicated by number, dorsal to ventral. D: distal tier. P: proximal tier. Species are coded by habitat depth, with species inhabiting greater depths indicated by increasingly dark symbol shades. (C) Sensitivity maxima in the same set of species, accounting for the effects of tiering and the presence of filter pigments in receptors in the second and third rows of the equatorial band. Otherwise coded as in panel B.

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Thomas W. Cronin and Kathryn D. Feller for the same species sets used for the visual pigment plots in Fig. 9.5B. Now a striking pattern is visible—in almost every receptor class, tuning shifts the sensitivity maximum toward spectral positions near the middle of the range (near ~500  nm) as habitat depth increases. Thus, shallow-living species have receptors with peak spectral sensitivities that can range from 400 nm to 700 nm, but the species inhabiting the deepest habitats have spectral ranges nearly 100 nm less. The selection of filters shapes the overall spectral sensitivity range of a species to occupy the expected spectral range of photons arriving at the depths it inhabits. The stomatopods exemplify how entire sets of photoreceptors can be adapted for different photic environments. There is an abundant and growing literature that explains how changes at specific amino acid sites in the opsin proteins of vertebrate visual pigments are involved in spectral tuning. In the visual pigments of marine vertebrates, both fishes and mammals, amino acid substitutions produce predictable shifts in visual pigment absorbance (see Douglas et al. 2003). Similarly, the molecular mechanisms producing spectral tuning in some mammalian cone visual pigments are becoming clear. Unfortunately, there is very little current understanding of the molecular adaptations that affect crustacean visual pigment tuning and function. The research results that do exist suggest that the mechanisms used will be unlike those of vertebrates. The deployment of visual pigments in crustacean photoreceptors also appears to be unusual compared with vertebrates, which is likely to be significant regarding the visual ecology of photoreception. Sakamoto et al. (1996) found that main rhabdoms of a grapsid crab, Hemigrapsus sanguineus, contain not one but two different (but closely related) opsin proteins. More recently, Rajkumar et al. (2010) discovered a similar system in a fiddler crab, Uca pugilator. Crab visual pigments are unusual in many ways, and they are distantly related to other crustacean (or other arthropod) types (see Cronin and Porter 2008). The duplicated expression pattern is not limited to crabs, however. The deep-sea mysid Gnathophausia ingens also has two closely related opsins in its main rhabdoms (Frank et al. 2009). The stomatopods, already proven to possess over a dozen spectral classes of photoreceptors, express a superabundance of visual opsins. In a given species, six opsin types are expected to form middle-wavelength visual pigments (four classes in main rhabdoms of the second and third rows of the midband, one class in the ventral two rows, and an additional one in the periphery), but twelve or more opsin sequences are expressed in some species (Porter et  al. 2009, Cronin et  al. 2010). How these are distributed among receptor cells is unknown.

SIGNAL PROCESSING AND CRUSTACEAN VISUAL ECOLOGY In ­chapter 8 in this volume, Glantz discusses how crustaceans process receptor signals to provide color vision and the ability to analyze polarized light. Here, we examine from an ecological perspective how systems of color and polarization vision are adaptive. Vision must of course be useful in the habitats and at the times when animals are active. The photic features that the animal experiences thus define the envelope within which visual spectral and polarization sensitivity can evolve. However, once a visual system comes into being, there is a second level of evolution that becomes available: its properties can shape the types of signals to which an animal can respond. In fact, the phenotypic reach of the visual system extends beyond the species housing it, because the signals of other animals—whether these are obvious, aposematic signals meant to advertise unpalatability or danger, or “negative signals” associated with camouflage of predators or prey—evolve within its capacities. In this way, environmental constraints on sensory systems also mold the evolution of crustacean phenotypes, as well as the signals of their predators and prey.



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The Visual Ecology of Crustacean Color Vision The presence of multiple classes of photoreceptors does not necessarily imply that color vision exists in a species. Daphnia magna, for example, has a small compound eye with four receptor types but displays simple reflexive responses (“color dances”) to different spectral stimuli (Smith and Baylor 1953, Smith and Macagno 1990, Cronin and Hariyama 2002, Marshall et al. 2003). Phototactic and photokinetic responses of planktonic crustaceans could easily involve similar reflexive spectral responses. Of course, these responses are adaptive and are valid research topics within visual ecology. Another example of the use of multiple spectral classes for tasks that do not involve color vision is seen in the beach amphipod Talorchestia longicornis. This animal uses a short-wavelength receptor class (peaking near 425 nm) for celestial orientation and one sensitive to longer wavelengths (~520 nm) for entrainment of circadian rhythms (Cohen et al. 2010). There is no current evidence that information from these classes is combined for analysis of color. In typical crustacean ommatidia, the R8 rhabdomere, containing a short-wavelength-sensitive visual pigment, sits above the fused rhabdom formed from rhabdomeres of R1-R7, which is middle-wavelength-sensitive (Martin and Mote 1982, Cronin and Hariyama 2002, Marshall et  al. 2003). The arrangement probably permits dichromatic (i.e., two-channel) color vision, and below we review some behavioral evidence that dichromatic crabs can discriminate color. However, no physiological exploration of the mechanisms used for color processing yet exists for any crustacean species. Most deep-sea species of crustaceans have, quite reasonably, become visual monochromats, dispensing with the R8 cell. Their main rhabdoms contain visual pigments adapted to detect downwelling light, bioluminescence, or hot water (Fig. 9.4). Mesopelagic oplophorid shrimp are an exception; some have two-tiered rhabdoms with the usual paired short/middle visual pigment classes (Cronin and Frank 1996). The relative size of the R8 component varies among species, decreasing with depth (Gaten et al. 2003), implying that species that frequently venture up into lighted waters may be able to process color, perhaps for prey detection or recognition of bioluminescence. Except for these deep-sea creatures, decapod crustaceans appear universally to be dichromats. Essentially all work on decapod color vision has centered on the crabs. At this point, no research has touched directly on the spectral adaptations of dichromatic crustaceans to natural scenes, but Chiao et al. (2000b) showed that the visual pigments of dichromatic fish, which tend to involve short-wavelength sensitivity maxima around 440 nm paired with middle-wavelength receptors peaking at 500–550 nm, are well suited to coral reef environments. These pairings are strikingly similar to those of the few coastal crab species that have been measured (e.g., Martin and Mote 1982, Cronin and Forward 1988), where R8 cells peak near 440 nm, while cells of the main rhabdom are maximally sensitive at about 510 nm, so crabs (at least) have reached the same receptor set solutions as fish in similar habitats. Color vision in crabs and its connection with intraspecific signaling is well documented. The charismatic little fiddler crabs, with their bright colors and their obvious use of motion signals involving the colorful major claw, are proven dichromats. Their patterns of visual pigment expression are like those of other crabs (Rajkumar et al. 2010). Fiddler crab colors vary among habitats. In the Australian species Uca vomeris, individual crabs change their body color to become more cryptic when predation pressure increases (Hemmi et al. 2006). It is not known whether this is visually monitored, however. More certain is the relationship between color vision and color signals in fiddler crabs. For instance, females of the Australian fiddler crab Uca mjoebergi use color vision to recognize mates (Detto 2007). Fig. 9.6A (see color version in center plate) illustrates how the dichromatic visual system of U. pugilator could visualize

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Fig. 9.6. Images of various crustaceans to show regions of the body used for signaling. (A) Male fiddler crab, Uca pugnax. The left panel is a standard black-and-white image, showing the achromatic appearance of this animal to the human eye. The right panel is a black-and white image derived from a color isoluminant image (see color insert in this volume). Here, each pixel in the color image has an identical R + G + B sum (255 bits), which is weighted by the photon catch for each of the two receptor classes of this species (the long-wavelength receptor’s input is printed with identical R and G values, to give yellow; the short-wavelength’s photon catch is printed in B). This derived black-and-white image shows parts of the crab that primarily stimulate the long-wavelength receptor as shades brighter than neutral; gray shades (i.e., the background) as neutral gray, and parts that primarily stimulate the short-wavelength receptor as shades darker than neutral. Note that much of the crab (particularly the claws and legs) is visible to the long-wavelength component of the crab’s visual system, contained in retinular cells R1-R8. (B) The single left panel shows a lateral view of the mantis shrimp Gonodactylus smithii. The four panels to the right show this same view, processed by pairs of receptors in the four tiered rows in the equatorial band of ommatidia, indicating how the visual system might process color information in the animal’s body coloration (see Chiao et al. 2000a for methods). The white and black arrows in the panel “Row 2” indicate color regions thought to serve as intraspecific signals. (C) Polarization signals seen on the keel (a structure projecting upward from the telson) of the stomatopod Odontodactylus cultrifer. The left set of images shows the keel from the right and left sides of the animal, photographed through a linear polarizer with the transmission axis of the polarizer indicated by the two-headed arrows. Note that the keel transmits vertically polarized light far more effectively than horizontally polarized light. The right set of images shows the same structure, from right and left sides, imaged through filters transmitting right (R) and left (L) circularly polarized light. Note that when viewed from the animal’s left side, the keel transmits left circular polarization more effectively, while the situation is reversed on the right side. Since these animals can see and analyze circular polarization, it is thought that this structure is used in cryptic signaling (see Chiou et al. 2008).



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important features of male crabs, particularly the claws and patches on the legs (which are sometimes raised during courtship). The most recent example of a role for color vision in mate choice involves evaluation of females by male crabs, and it comes from an entirely different crab family, the portunid Callinectes sapidus. Baldwin and Johnsen (2009) found that male blue crabs prefer females with red dactyls on their claws, and that the choice is color, not brightness, based. One final case of decapod crustacean vision potentially associated with sexual signaling is not from a crab at all, but instead involves the freshwater crayfish Procambarus clarkii. These animals express seasonal changes in their visual pigments, and they are more sexually active in the winter. They are also sexually dimorphic, and it is possible that the more complex set of receptors present during the winter is involved in mate choice (Cronin and Hariyama 2002). Finally, crayfish can also remember and visually recognize conspecifics they have met before, an ability that certainly would be enhanced if color vision is involved (Van der Velden et al. 2008). Finally, we move to the polychromatic stomatopods. Their complex retinas are seemingly the ultimate in color vision design, but their odd sampling of space, in which the set of receptors that contain the color apparatus views only a planar slice through space (e.g., Fig. 9.2D), implies that registering the spatial location of color (and possibly polarization) within the extended visual fields of the peripheral sets of ommatidia could be difficult. It is thought that small scanning movements made independently by the two eyes are involved in coordinating chromatic, polarizational, and spatial vision in these animals (Land et al. 1990). Stomatopods can be trained to discriminate among colored objects, the standard test for color vision (Marshall et al. 1996, Cronin and Marshall 2004)—in fact, they are the only crustaceans that have been tested in purely color-discrimination tasks independent of other biological cues (the examples described above are strictly related to mate selection). The ecological role of color vision is not proven, but with their bright colors and contrasting markings, it seems certain that its functions include intraspecific signaling (Caldwell and Dingle 1975). Note that color patches on the bodies of stomatopods could be visually enhanced considerably by processing within sets of color receptors known to be present in these animals (Fig. 9.6B). The Visual Ecology of Crustacean Polarization Vision Nearly all crustaceans have receptors potentially capable of analyzing polarized light (see ­chapter 9 in this volume). Even many deep-sea crustaceans, including euphausiids and hyperiid amphipods (e.g., Meyer-Rochow 1978, Meyer-Rochow and Walsh 1978, Hallberg and Nilsson 1983) have rhabdoms with the characteristic crustacean pattern of alternating, orthogonal layers of microvilli; although as expected, the deepest-living species, especially the vent inhabitants, lack this organization (Gaten et al. 1992, O’Neill et al. 1995, Jinks et al. 2002). Obviously, not every species with alternating microvillar layers has been assessed for polarization sensitivity, but it is reasonable to assume that the high degree of organization is associated with this capacity, and no counterexamples have been reported. The classic review of polarization vision in invertebrates, with many crustacean examples, is that of Waterman (1981). Important, more recent reviews are those of Horváth and Varjú (2004) and Marshall et al. (2011). The physiology and visual ecology of polarized-light sensitivity is far better investigated in insects than in crustaceans, but crustacean examples nevertheless abound. As in insects, the polarized-light pattern of the sky permits orientation of some crustaceans living on beaches (reviewed in Waterman 1981). Fiddler crabs have richly detailed worlds of polarized light, and it is certainly possible that they use such cues for short-range orientation, perhaps when returning to the burrow upon being threatened (Zeil and Hemmi 2006). Even

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Thomas W. Cronin and Kathryn D. Feller shallow-water crustaceans, such as the cladoceran Daphnia, will orient in polarized light fields (see Waterman 1981). Although the literature is rich in studies of spontaneous polarization orientation in crustaceans, there are few clear examples of a role for polarization sensitivity in crustacean ecology and behavior. One exception involves the grass shrimp, Palaemonetes vulgaris, which orients its escape responses in shallow water using the transmitted skylight polarization pattern (Goddard and Forward 1991). Polarization vision, as opposed to polarization sensitivity, is a sensory function analogous to color vision, giving an animal the ability to recognize features of a visual scene and to discriminate objects based solely on their differences in polarization. Humans, of course, lack a refined ability to see polarization and are completely unaware of its variation in scenes, so it is difficult for us to appreciate how common this type of vision is among animals. Most crustaceans, on the other hand, are probably aware of polarized light’s distribution. It is likely to play a more important role in their lives than color does; in fact, the neuroanatomy that underlies color vision in mantis shrimp is derived from an earlier polarization analytical system (Marshall et al. 2008). Polarized-light vision is often suspected of being important in contrast enhancement, particularly in contrast-poor aquatic habitats (Johnsen et al. 2011, Marshall and Cronin 2011), but there is no convincing evidence that crustaceans enhance contrast using their polarization sense. What is known, however, is that members of at least one crustacean group, the stomatopods, have true polarization vision (Marshall et al. 1999). Stomatopods readily learn to discriminate among objects using only polarization differences. This ability certainly could be useful for predatory or orientation behavior. It is also associated with signaling in many stomatopod species (Cronin et al. 2003), which display linearly polarized markings and appendages during courtship behavior (Chiou et al. 2011) and in agonistic interactions (Fig. 9.6C). The optics of the polarization reflectors are often exotic and in some cases, not well understood. A few species of mantis shrimp are actually thought to communicate using circularly polarized light, instead of the more commonly perceived linearly polarized type (Chiou et al. 2008; see Fig. 9.6C). Such signals would be conspicuous and unambiguous in the weakly polarized surroundings of marine habitats. Polarization signals in general are most useful in crustaceans that live away from surface waters, where color is the more reliable signal. They are likely to be more common than is realized among crustaceans inhabiting moderate depths. The circular polarization receptors that allow stomatopods to recognize their conspecifics are based on optics that spectrally outperform any artificial devices ever built (Roberts et al. 2009).

VISUAL DEVELOPMENT AND CRUSTACEAN VISUAL ECOLOGY The Different Visual Worlds of Larval and Adult Crustaceans Many crustaceans have life cycles in which the early stages are adapted to occupy a dramatically different environment than the adult. Their larvae live in the pelagic environment as transient members of the plankton before metamorphosing into benthic or pelagic adults. These animals are often referred to as meroplanktonic, since they are only planktonic over one phase of their life history, the larval period. The separation of the larval and adult niches serves to distribute the species within a geographic area as well as reduce resource competition and inbreeding within a single habitat (Pechenik 1999). Occupying a spectrally distinct habitat, larvae have evolved a suite of behaviors and adaptations that are specific to the ecology of planktonic life. Thus, larval eyes are commonly quite different from adult eyes in both form and function. Consider the hydrothermal vent crab B.  thermydron, introduced earlier for its naked retina



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adapted to detect infrared light emitted by hot water. Despite the unusual adult eye design and function, the pelagic larvae of this species possess compound eyes like those of other crab larvae. Furthermore, as described later, these eyes are spectrally adapted for living in a blue, open water habitat and not for viewing infrared light at all (Jinks et al. 2002). This extreme example documents the adaptive changes that can take place in a crustacean visual system over an individual’s life history. In this section, we explore current knowledge regarding larval visual adaptations to their photic environments. The field of crustacean larval sensory biology is relatively new; consequently, larval visual ecology remains an underdeveloped field for the majority of crustacean species. Here, we primarily refer to compound eyes of larval malacostracan crustaceans. We specifically focus on decapods, stomatopods, and euphausiids, though other crustaceans will be mentioned as available. Larval Eyes: Relating Evolution and Development Studies of larval visual systems are particularly valuable for the evolutionary information they yield. Transparent apposition compound eyes are of course beneficial for camouflage in the pelagic environment, but the design also holds the key to understanding compound eye evolution. Apposition-type eyes are thought to be the evolutionary ancestors of superposition eyes, not because larvae possess apposition optics, but rather because the transparent apposition eye is preadapted to form superposition optics (Nilsson 1983). Larval eyes necessarily form superposition rays as a secondary effect of their elongated crystalline cones. Thus, when species with adult superposition eyes undergo metamorphosis, the existing larval tissue is preadapted to be remodeled into refracting or reflecting superposition eyes (Palaemonetes: Douglass and Forward 1989; Penaeus: Elofsson 1969; Panulirus: Meyer-Rochow 1975; Palaemon: Fincham 1984; Neomysis and Thysanoessa: Nilsson et al. 1986; Oplophorus: Gaten and Herring 1995). The optical properties of most transparent larval eyes provide an opening for the evolution of the modern diversity of crustacean compound eye designs. The only known exception is the euphausiid Thysanopoda tricuspidata, which possesses a refracting superposition larval eye. The evolutionary implications of studies of larval visual systems extend beyond the ontogeny of eye structure. Research on eye development provides characters that help resolve the often confusing phylogenetic relationships among arthropod species (Harzsch 2006, Fischer and Scholtz 2010). For instance, studies of the pattern of cellular development in the eyes of lobsters, crabs, and shrimp reveal a high degree of homology between these crustaceans and insects (Harzsch et al. 1999). This is not surprising, since (as noted earlier) insects are phylogenetically nested within the crustaceans. Eyes of both larval and adult crustaceans are the product of millions of years of selection. Therefore, research on the development, structure, and function of the eyes of species that undergo metamorphosis not only provides insight into the larger picture of arthropod eye evolution but also reveals how animal eyes adapt and evolve to changing environments and behaviors, even within a single organism (for further discussion, see Dangles et al. 2009). The Ontogeny of Visual Pigment Expression Earlier, the ecology of visual pigments was discussed at length for adult crustaceans living in different aquatic environments. What better model could be imagined for examining spectral adaptation than the sequential forms of a single animal adapted to live in disparate habitats over its lifetime! Presumably, animals that spend part of their life in one type of light environment, but then migrate to another, will possess visual pigments adapted to each life phase. Changes

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Thomas W. Cronin and Kathryn D. Feller in visual pigment expression have been documented in many aquatic vertebrates that occupy different photic environments over the course of their ontogeny. In contrast, only a handful of studies have considered how visual pigments vary in different environments over the ontogeny of crustaceans, or indeed, any marine invertebrates. In one example (Jinks et al. 2002), zoeal larvae of the hydrothermal vent crab, B. thermydron, express a visual pigment with a λ max of 447 nm, well suited for detecting the blue light of the mesopelagic environment. As the animal migrates toward its adult home near deep-sea hydrothermal vents, each successive stage of development expresses an increasingly red-shifted visual pigment (megalopae λ max 479 nm; adult λ max 489 nm). Whether the ontogenetic changes in photoreceptor sensitivity seen in B. thermydron are common among brachyuran crabs or other crustaceans is yet to be elucidated. The only other crab that has been investigated for ontogenetic changes in visual pigment expression is the blue crab, C. sapidus. Photoreceptors in megalopae and adults of this species have similar spectral absorbances (λ max = 504 nm; Cronin et al. 1995). It is not surprising that megalopal and adult blue crabs have the same visual pigment, since they are both found in similar shallow marine and estuarine habitats. The zoeal larvae, however, live in the bluer light environment of the open ocean. Given the ontogenetic change in light environment, it is reasonable to predict that blue crab larvae may express a visual pigment shifted to view the shorter wavelengths that dominate the habitat in which they reside. However, until spectral absorption data are collected, the nature of the larval blue crab visual pigments as they relate to the adult stage remains conjectural. In accordance with everything bizarre and unusual about stomatopod eyes, the ontogeny of the retina and the changing expression of visual pigments in these seemingly alien crustaceans is no exception. As previously mentioned, adult stomatopod crustaceans can possess a dozen or more distinct visual receptor classes. In the eyes of four species of stomatopod larvae that have been investigated, however, evidence for only one spectral class has been found, which is reasonably well matched to their blue pelagic environments. However, two of these are Pullosquilla larval species (P. thomassini and P. littoralis; Jutte et al. 1998), which have visual pigments with extremely short λ max values (446 nm). This may be a result of phylogenetic constraint in these species, or the unusually short-wavelength absorbance may reflect an unrecognized feature of the larval ecology. It is unclear whether larval visual pigments are expressed only in the larval retina or if they are re-expressed in receptors of the adult eye after metamorphosis. One thing is for certain: in stomatopods, the larval retina itself is separate from the adult’s. Rather than modify the existing larval retina into the adult morphotype, as typical malacostracans do, stomatopods actually form an adult retina adjacent to and separate from the larval structure (Fig. 9.3A, C). The result: two retinas in a single eye! It is therefore possible that stomatopods, which have completely separate larval and adult lifestyles, environments, and retinal tissue, may also possess completely separate visual pigments in their successive retinas. The evidence is not clear and may even support the opposite situation in some cases. For instance, larvae of Squilla empusa, a stomatopod with only one known adult visual pigment (λ max 507 nm), express a larval pigment (λ max 509 nm) similar or identical to the adult’s (Cronin and Jinks 2001). These data suggest that, despite the development of separate retinas during metamorphosis, the visual pigment may be conserved throughout the lifetime of S. empusa. Similarly, in the two species of Pullosquilla larvae that possess receptors with λ max values ~446 nm, the corresponding adult retinas have a class of photoreceptors with nearly identical spectral absorption (Jutte et al. 1998). While the double retina is certainly an unusual eye developmental mechanism, it is not entirely unique to stomatopods. The mesopelagic euphausiid T. tricuspidata also develops its larval retina as a separate structure from the adult eye. During the furcilia or later stage larva, an eye composed of seven crystalline cones and approximately 90 rhabdoms appears. This eye is atypical since it uses image-forming superposition optics rather than the transparent apposition



Sensory Ecology of Vision in Crustaceans

optics usually found in the eyes of crustacean larvae (Land 1981). In a similar fashion to the stomatopods, T. tricuspidata develops a new retina along the anterior portion of the existing eye during its late stage larval phase that will become the adult eye. This eye grows and extends posteriorly until the larval eye is completely engulfed, eventually ejecting the original seven crystalline cones during the late adolescent stage (Land 1981). Like the stomatopods, T. tricuspidata (and likely other euphausiids) provides an excellent model for examining visual adaptations of separately evolved eyes within a single organism. Expansion of visual pigment datasets for both stomatopods and T. triscupidata will provide insight to the mechanisms that drive visual pigment expression pattern evolution (e.g., phylogenetic or developmental constraints, differences in adult settlement habitats, or simply differences in ecology). As shown here, larval crustacean visual ecology is a very underdeveloped field relative to the corresponding research that exists for adult crustaceans and other marine animals. Not only do we lack understanding of the visual pigments in the main rhabdoms of most larvae but also we have yet to confirm the presence of ultraviolet-sensitive R8 cells in their retinas. R8 cells have been morphologically identified in the larval eyes of shrimp (Palaemonetes pugio, Douglass and Forward 1989); however, it is unknown what visual pigments reside in these cells. Though some behavioral studies show that crab larvae respond to ultraviolet light (Forward and Cronin 1979), such responses could be based on either ultraviolet-sensitive visual pigments or an extended absorption by the middle-wavelength visual pigments of the main rhabdom. Larval Visual Ecology The mere presence of sophisticated compound eyes in crustacean larvae suggests a prominent role for relatively complex visually mediated behaviors. Larvae are not simply low-tech versions of adults; they are exquisitely adapted miniature crustaceans. Larvae use their visual systems to hunt for food, maintain proper orientation in the water column, locate the adult habitat, and avoid planktonic predators such as ctenophores and coelenterate medusae in estuaries, or juvenile yellowfin tuna in the open ocean (Bullard et al. 1999, Graham et al. 2007, Forward 2009). Many larvae display a strong visually mediated predator avoidance response, known as the shadow response. A sudden decrease in light intensity elicits a rapid escape response whereby the animals actively swim or passively sink away from the surface (reviewed in Forward 2009). In addition to predation and predator avoidance, larvae must also find an appropriate adult habitat in which to settle and reside as an adult. Crustacean larvae often settle and metamorphose in a place they have not previously experienced but that is nonetheless suitable to promote survival. They are genetically programmed to find the right location in which the adult stage is adapted to live. While this complex behavior requires the input of several sensory systems, visual cues appear to be important. When presented with a choice, a variety of reef crustacean species will select the appropriate settlement habitat based on a combination of olfactory and visual cues (Lecchini et  al. 2010). The specific visual cues required for habitat selection are unknown, since they have yet to be examined independently of olfactory stimuli. Differences in spectral sensitivities among larval species may be associated with adult habitat-seeking behavior; however, at present this must remain merely a hypothesis. A fundamental light-mediated behavior of many marine zooplankton is their daily migration toward and away from the surface, or diel vertical migration. With each setting and rising of the sun, larvae of many crustacean species take their cue to respectively ascend and descend in the water column. Diel vertical migrations allow animals to avoid visually oriented surface predators (Forward and Hettler 1992) as well as potentially damaging ultraviolet radiation (Lampert 1989, Leech and Johnsen 2003) by traveling to deeper, darker depths during the day, returning to feed near the surface at night (reviewed in Cohen and Forward 2009). The requirement for

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Thomas W. Cronin and Kathryn D. Feller well-controlled vertical migrations implies that spectral sensitivity functions of larval crustacean photoreceptors could be tuned for best function at the times of migration. Vertical migrations of planktonic crustaceans, including crab larvae, are partially governed by endogenous circadian and circatidal rhythms (Enright and Hamner 1967, Cronin and Forward 1979). Setting these rhythms involves (at least in part) photoperiodic cues, which in most animals are handled by extraocular, nonvisual photoreceptors. In certain cases visual photoreceptors may also play a role in photoentrainment, as shown by recent photobehavior studies on the amphipod T. longicornis (Cohen et al. 2010). While planktonic larvae must respond to light, true compound eyes need not be present for photodependent behaviors to occur. Many pelagic larvae, including the earliest stages of copepods, branchiopods (e.g., Artemia), barnacles (Hallberg and Elofsson 1983), and decapod shrimps (Elofsson 1969), possess a cluster of pigmented light-sensitive cells known as a nauplius eye. The nauplius eye arises from a medial anlage of a few densely packed ocelli and lacks the optics and ordered structure of a compound eye. Despite the paucity of spatial information that such an eye provides, larvae with nauplius eyes carry out well-oriented phototactic behaviors, moving toward or away from a source of light. Given that the ecological demands placed on an animal shape the evolution of its sensory structures, larvae with true compound eyes presumably must perform more complex visual tasks than those that express only a nauplius eye. Barnacles provide an interesting example of how the changing ecological needs of an animal are reflected in the ontogeny of eye structure, particularly regarding changes in ocular complexity. As sessile animals, adult barnacles have little need for a complex compound eye. They get by with a simple light-sensing structure, essentially a degenerate form of a compound eye present in the previous stage, the semipelagic cyprid larva. The earliest larval stages, themselves called nauplii, have eponymous nauplius eyes. These are followed by the cyprid, whose main goal is to locate a suitable surface on which to attach and metamorphose into an adult. The requirements of such a relatively complex task have led to the formation of a true, if rather simple, compound eye, present only during the cyprid stage (Hallberg and Elofsson 1983). Barnacles are thus among the many species that exemplify how ontogenetic changes in ecology can shape the structure and function of crustacean eyes.

FUTURE DIRECTIONS The sensory ecology of crustacean vision is not a field that lacks research questions! As is plain from the preceding sections, there are a number of problems that have been studied only rarely, or not at all. On the other hand, the field is benefiting from the rapid development of new tools and approaches that open up access to research projects too difficult to approach only a few years ago. These tools range from ones involved with field research, such as underwater photographic and video tools and new generations of miniature or submersible spectrometers, down to effective methods for dealing with gene sequences, expression patterns, and evolution of the molecules involved in photoreception and ocular development. Here, we briefly discuss research areas that we perceive to be open avenues for progress in the near future. For well over a century, studies of the optics of crustacean eyes have uncovered novel mechanisms of light handling, new to science and unlike previous manmade optics (e.g., Exner 1891, Land 1984, Nilsson 1989). These have involved image forming systems, such as reflecting superposition (which has found application in x-ray telescopes; Putkunz and Peele 2009), and photonic structures that manipulate light’s state of polarization (Roberts et  al. 2009). Similarly, crustaceans reveal unexpected mechanisms of color analysis (Cronin and Marshall 1989) and visual pigment organization (Sakamoto et al. 1986). Any research on crustacean ocular anatomy



Sensory Ecology of Vision in Crustaceans

or visual function has the potential to open new doors—on occasion, doors that were not even known to be closed—something that sensory biologists should always keep in mind. Who could have foreseen that eyes completely lacking optics, incorporating blue-sensitive visual pigments, would be used to detect outflow from thermal vents, for instance? Research on the structure-function relationships of crustacean visual pigments is still in its earliest phases. These proteins, while evolutionarily conserved and obviously related to the visual pigments of vertebrates, use poorly understood mechanisms of spectral tuning and of interacting with each other and with molecules in photoreceptor cells. Multiple expression of visual pigments in crustacean photoreceptors seems to be quite common, perhaps even the rule, but no functional explanation for this has been found. A related topic is the diversity, function, and distribution of extraocular opsins in crustaceans. Other than studies of vision, very little is known of how these animals sense light. New research on the sensory ecology of nonvisual photoreception offers promise of better understanding of crustacean photobiology and of the role of extraocular photoreception in other animals. Neural processing of visual signals is much less researched in crustaceans than in insects, despite the highly homologous (and similarly organized) neuroanatomy. New mechanisms of observing coordinated activity in groups of neurons, and of examining central nervous structures in vivo or at least in situ, provide promising avenues for research. Both color and polarization carry information that crustaceans use in novel situations, and new research here is certain to be beneficial for both basic understanding of crustacean sensory biology and ecology and for applications to artificial imaging and image-analysis systems. Finally, this chapter has raised a number of questions regarding visual development in crustaceans and has noted the scarcity of information about larval visual ecology in these animals. To be fair, the same could be said about studies of plankton and larvae in general, not just larval—or indeed, planktonic—crustaceans. The developmental sequences of optics are well studied, but the physiology of larval eyes and their corresponding adult forms, particularly in crustaceans such as stomatopods, where the entire retina is replaced, is barely investigated at all. Larvae are fully adapted crustaceans and should never be viewed as temporarily patched-together way stations on the route to making a properly built adult.

SUMMARY AND CONCLUSIONS Returning to the opening message of this chapter, crustaceans sport the most diverse visual systems of any animal group, and they inhabit nearly the full range of habitats occupied by life on earth. Their vision is highly adapted, often exotic, and in many species offers their primary sensory contact with the external world. Studies of the sensory ecology of crustacean vision are worth doing in their own right, and they often offer astonishing insights into how animals meet and cope with the environmental challenges they must face. Whether the level of investigation is at the level of the gene, or of behavior, or of the features of the environment occupied by the crustacean species of interest, entering the sensory worlds of crustaceans rarely fails to be exciting and rewarding.

REFERENCES Ala-Laurila, P., K. Donner, and A. Koskelainen. 2004. Thermal activation and photoactivation of visual pigments. Biophysical Journal 86:3653–3662.

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Thomas W. Cronin and Kathryn D. Feller Baldwin, J., and S. Johnsen. 2009. The importance of color in mate choice of the blue crab Callinectes sapidus. Journal of Experimental Biology 212:3762–3768. Bullard, S.G., N.L. Lindquist, and M.E. Hay. 1999. Susceptibility of invertebrate larvae to predators: how common are postcapture larval defenses? Marine Ecology Progress Series 191:153–161. Caldwell, R.L., and H. Dingle. 1975. Ecology and evolution of agonistic behavior in stomatopods. Naturwissenschaften 62:214–222. Chiao, C.C., T.W. Cronin, and N.J. Marshall. 2000a. Eye design and color signaling in a stomatopod crustacean, Gonodactylus smithii. Brain, Behavior and Evolution 56:107–122. Chiao, C.C., M. Vorobyev, T.W. Cronin, and D. Osorio. 2000b. Spectral tuning of dichromats to natural scenes. Vision Research 40:3257–3271. Chiou, T.-H., S. Kleinlogel, T. Cronin, R. Caldwell, B. Lofle, A. Siddiqi, A. Goldizen, and J. Marshall. 2008. Circular polarisation vision in a stomatopod crustacean. Current Biology 18:429–434. Chiou, T.-H., N.J. Marshall, R.L. Caldwell, and T.W. Cronin. 2011. Changes in light reflecting properties of signaling appendages alter mate choice behaviour in a stomatopod crustacean Haptosquilla trispinosa. Marine and Freshwater Physiology and Behaviour 44:1–11. Chamberlain, S.C. 2000. Vision in hydrothermal vent shrimp. Philosophical Transactions of the Royal Society of London B – Biological Sciences 355:1151–1154. Cohen, J.H., and R.B. Forward, Jr. 2009. Zooplankton diel vertical migration: a review of proximate control. Pages 77–110 in R. Gibson, R. Atkinson, and J. Gordon, editors. Oceanography and marine biology: an annual review. Vol. 48. CRC Press, New York. Cohen, J.H., T.W. Cronin, N. Lessios, and R.B. Forward, Jr. 2010. Visual physiology underlying orientation and diel behavior in the sand beach amphipod Talorchestia longicornis. Journal of Experimental Biology 213:3843–3851. Cronin, T.W. 1986. Optical design and evolutionary adaptation in crustacean compound eyes. Journal of Crustacean Biology 6:1–23. Cronin, T.W. 2008. Visual ecology. Pages 211–246 in R. Masland and T.D. Albright, editors. The senses: a comprehensive reference, Vol. 1, Vision. Academic Press, San Diego. Cronin, T.W., and R.B. Forward, Jr. 1979. Tidal vertical migration: an endogenous rhythm in estuarine crab larvae. Science 205:1020–1022. Cronin, T.W., and R.B. Forward, Jr. 1988. The visual pigments of crabs. I. Spectral characteristics. Journal of Comparative Physiology A 162:463–478. Cronin, T.W., and T.M. Frank. 1996. A short-wavelength photoreceptor class in a deep-sea shrimp. Proceedings of the Royal Society of London B – Biological Sciences 263:861–865. Cronin, T.W., and T. Hariyama. 2002. Spectral sensitivity in crustaceans. Pages 499–511 in K. Wiese, editor. The crustacean nervous system. Springer, Berlin Heidelberg New York. Cronin, T.W., and R.N. Jinks. 2001. Ontogeny of vision in marine crustaceans. American Zoologist 41:1098–1108. Cronin, T.W., and N.J. Marshall. 1989. A retina with at least ten spectral types of photoreceptors in a stomatopod crustacean. Nature 339:137–140. Cronin, T.W., and N.J. Marshall. 2004. The unique visual world of mantis shrimps. Pages 39–268 in F. Prete, editor. Complex worlds from simpler nervous systems. MIT Press, Cambridge, MA. Cronin, T.W., and M.L. Porter. 2008. Exceptional variation on a common theme: the evolution of crustacean compound eyes. Evolution: Education and Outreach 1:463–475. Cronin, T.W., N.J. Marshall, and R.L. Caldwell. 2000. Spectral tuning and the visual ecology of mantis shrimps. Philosophical Transactions of the Royal Society of London B – Biological Sciences 355:1263–1268. Cronin, T.W., N.J. Marshall, R.L. Caldwell, and D. Pales. 1995. Compound eyes and ocular pigmets of crustacean larvae (Stomatopoda and Decapoda, Brachyura). Marine Behaviour and Physiology 26:219–231. Cronin, T.W., N. Shashar, R.L. Caldwell, N.J. Marshall, A.G. Cheroske, and T.H. Chiou. 2003. Polarization vision and its role in biological signaling. Integrative and Comparative Biology 43:549–558.



Sensory Ecology of Vision in Crustaceans

Cronin, T.W., M.L. Porter, M.J. Bok, J.B. Wolf, and P.R. Robinson. 2010. The molecular genetics and evolution of colour and polarization vision in stomatopod crustaceans. Ophthalmic and Physiological Optics 30:460–469. Dangles, O., D. Irschick, L. Chittka, and J. Casas. 2009. Variability in sensory ecology: expanding the bridge between physiology and evolutionary biology. Quarterly Review of Biology 84:51–74. Detto, T. 2007. The fiddler crab Uca mjoebergi uses colour vision in mate choice. Proceedings of the Royal Society of London B—Biological Sciences 274:2785–2790. Douglas, R.H., D.M. Hunt, and J.K. Bowmaker. 2003. Spectral sensitivity tuning in the deep-sea. Pages 323–342 in S.C. Collin and N.J. Marshall, editors. Sensory processing in aquatic environments. Springer, New York. Douglass, J.K., and R.B. Forward, Jr. 1989. The ontogeny of facultative superposition optics in a shrimp eye: hatching through metamorphosis. Cell and Tissue Research 258:289–300. Elofsson, R. 1969. The development of the compound eyes of Penaeus duorarum (Crustacea: Decapoda) with remarks on the nervous system. Cell and Tissue Research 97:323–350. Enright, J.T., and W.N. Hamner. 1968. Vertical diurnal migration and endogenous rhythmicity. Science 157:937–941. Exner, S. 1891. The physiology of the compound eyes of insects and crustaceans. Translated from the German by R.C. Hardie (1989), republished by Springer, Heidelberg. Fincham, A.A. 1984. Ontogeny and optics of the eyes of the common prawn Palaemon (Palaemon) serratus (Pennant, 1777). Zoological Journal of the Linnean Society 81:89–113. Fischer, A., and G. Scholtz. 2010. Axogenesis in the stomatopod crustacean Gonodactylaceus falcatus (Malacostraca). Invertebrate Biology 129:59–76. Forward, R.B. Jr. 2009. Larval biology of the crab Rhithropanopeus harrisii (Gould): a synthesis. Biological Bulletin 216:243–256. Forward, R.B. Jr., and T.W. Cronin. 1979. Spectral sensitivity of larvae from intertidal crustaceans. Journal of Comparative Physiology A 133:311–315. Forward, R.B. Jr., and W.F. Hettler Jr. 1992. Effects of feeding and predator exposure on photoresponses during diel vertical migration of brine shrimp larvae. Limnology and Oceanography 37:1261–1270. Frank, T.M., M.L. Porter, and T.W. Cronin. 2009. Spectral sensitivity, visual pigments and screening pigments in two life history stages of the ontogenetic migrator Gnathophausia ingens. Journal of Marine Biological Association of the United Kingdom 89:119–129. Gaten, E., and P.J. Herring. 1995. Morphology of the reflecting superposition eyes of larval oplophorid shrimps. Journal of Morphology 225:19–29. Gaten, E., P.M.J. Shelton, and P.J. Herring. 1992. Regional morphological variation in the compound eyes of mesopelagic decapods in relation to their habitat. Journal of the Marine Biological Association of the United Kingdom 72:61–75. Gaten, E., P.J. Herring, and P.M.J. Shelton. 2002. Eye morphology and optics of the double-eyed mysid Euchaetomera typical. Acta Zoologica 83:221–230. Gaten, E., P.M.J. Shelton, and M.S. Nowel. 2003. Interspecific variations in the rhabdom structure of oplophorid shrimps. Journal of Morphology 257:87–95. Graham, B.S., D. Grubbs, K. Holland, and B.N. Popp. 2007. A rapid ontogenetic shift in the diet of juvenile yellowfin tuna from Hawaii. Marine Biology 150:647–658. Goddard, S., and R.B. Forward, Jr. 1991. The role of the underwater polarized light pattern in sun compass navigation of the grass shrimp, Palaemonetes vulgaris. Journal of Comparative Physiology A 169:479–491. Hallberg, E. 1978. The fine structure of the compound eyes of mysids (Crustacea: Mysidae). Cell and Tissue Research 184:45–65. Hallberg, E., and R. Elofsson. 1983. The larval compound eye of barnacles. Journal of Crustacean Biology 3:17–24. Hallberg, E., and D.-E. Nilsson. 1983. The euphausiid compound eye—a morphological re-investigation (Crustacea: Euphausiacea). Zoomorphology 103:59–66.

259

260

Thomas W. Cronin and Kathryn D. Feller Harzsch, S. 2006. Neurophylogeny: architecture of the nervous system and a fresh view on arthropod phyologeny. Integrative and Comparative Biology 46:162–194. Harzsch, S., J. Benton, R.R. Dawirs, and B. Beltz. 1999. A new look at embryonic development of the visual system in decapod crustaceans: neuropil formation, neurogenesis, and apoptotic cell death. Journal of Neurobiology 39:294–306. Hemmi, J.M., N.J. Marshall, W. Pix, M. Vorobyev, and J. Zeil. 2006. The variable colours of the fiddler crab Uca vomeris and their relation to background and predation. Journal of Experimental Biology 209:4140–4153. Horridge, G.A. 1978. The separation of visual axes in apposition compound eyes. Philosophical Transactions of the Royal Society of London B—Biological Sciences 285:1–59. Horváth, G., and D. Varjú. 2004. Polarized light in animal vision. Springer, Berlin Heidelberg. Jinks, R.N., T.L. Markley, E.E. Taylor, G. Perovich, A.I. Dittel, C.E. Epifanio, and T.W. Cronin. 2002. Adaptive visual metamorphosis in a deep-sea hydrothermal vent crab. Nature 420:68–70. Johnsen, S., N.J. Marshall, and E.A. Widder. 201l. Polarization sensitivity as a contrast enhancer in pelagic predators: lessons from in situ polarization imaging of transparent zooplankton. Philosophical Transactions of the Royal Society of London, B – Biological Sciences 366:655–670. Jordão, J.M., T.W. Cronin, and R.F. Oliveira. 2007. Spectral sensitivity of four species of fiddler crabs (Uca pugnax, Uca pugilator, Uca vomeris and Uca tangeri) measured by in situ microspectrophotometry. Journal of Experimental Biology 210:447–453. Jutte, P.A., T.W. Cronin, and R.L. Caldwell. 1998. Photoreception in the planktonic larvae of two species of Pullosquilla, a lysiosquilloid stomatopod crustacean. Journal of Experimental Biology 201:2481–2488. Lampert, W. 1989. The adaptive significance of diel vertical migration of zooplankton. Functional Ecology 3:21–28. Land, M.F. 1981. Optics and vision in invertebrates. Pages 471–592 in H. Autrum, editor. Comparative physiology and evolution of invertebrates B, Invertebrate visual centers and behavior, I. Handbook of sensory physiology VII/6B. Springer, Berlin. Land, M.F. 1984. Crustacea. Pages 401–438 in M.A. Ali, editor. Photoreception and vision in invertebrates. Plenum Press, New York. Land, M.F. 1988. The functions of eye and body movements in Labidocera and other copepods. Journal of Experimental Biology 140:381–391. Land, M.F. 1989. The eyes of hyperiid amphipods: relations of optical structure to depth. Journal of Comparative Physiology A 164:751–762. Land, M.F. 2000. On the functions of double eyes in midwater animals. Philosophical Transactions of the Royal Society of London B—Biological Sciences 355:1147–1150. Land, M.F. 2002. What is a naked retina good for? Nature 420:30–31. Land, M.F., and D.-E. Nilsson. 2002. Animal eyes. Oxford University Press, Oxford UK. Land, M.F., F.A. Burton, and V.B. Meyer-Rochow. 1979. The optical geometry of euphausiid eyes. Journal of Comparative Physiology A 130:49–62. Land, M.F., N.J. Marshall, D. Brownless, and T.W. Cronin. 1990. The eye-movements of the mantis shrimp Odontodactylus scyllarus (Crustacea: Stomatopoda). Journal of Comparative Physiology A 167:155–166. Lecchini, D., S.C. Mills, C. Brie, R. Maurin, and B. Banaigs. 2010. Ecological determinants and sensory mechanisms in habitat selection of crustacean postlarvae. Behavioral Ecology 21:599–608. Leech, D., and S. Johnsen. 2003. Behavioral responses—UVR avoidance and vision. Pages 455–481 in E. Helbling and H. Zagarese, editors. UV effects in aquatic organisms and ecosystems. The Royal Society of Chemistry, Cambridge. Marshall, N.J., and T.W. Cronin. 2011. Polarisation vision. Current Biology 21:R101–R105. Marshall, N.J., and M.F. Land. 1993. Some optical features of the eyes of stomatopods. I. Eye shape, optical axis, and resolution. Journal of Comparative Physiology A 173:565–582. Marshall, N.J., and J. Oberwinkler. 1999. The colourful world of the mantis shrimp. Nature 401:873–874. Marshall, N.J., J.P. Jones, and T.W. Cronin. 1996. Behavioural evidence for color vision in stomatopod crustaceans. Journal of Comparative Physiology A 179:473–481.



Sensory Ecology of Vision in Crustaceans

Marshall, N.J., T.W. Cronin, N. Shashar, and M.F. Land. 1999. Behavioural evidence for polarisation vision in stomatopods reveals a potential channel for communication. Current Biology 9:755–758. Marshall, N.J., T.W. Cronin, and T.M. Frank. 2003. Visual adaptations in crustaceans: chromatic, developmental, and temporal aspects. Pages 343–372 in S.C. Collin and N.J. Marshall, editors. Sensory processing in aquatic environments. Springer, New York. Marshall, N.J., T.W. Cronin, and S. Kleinlogel. 2008. A review of stomatopod eye structure and function. Arthropod Structure and Development 36:420–448. Marshall, N.J., T.W. Cronin, and M.F. Wehling. 2011. New directions in biological research (theme issue). Philosophical Transactions of the Royal Society of London B – Biological Sciences 366:611–782. Martin, F.G., and M.I. Mote. 1982. Color receptors in marine crustaceans: a second spectral class of retinular cell in the compound eyes of Callinectes and Carcinus. Journal of Comparative Physiology A 145:549–554. Meyer-Rochow, V.B. 1975. Larval and adult eye of the western rock lobster (Panulirus longipes). Cell and Tissue Research 162:439–458. Meyer-Rochow, V.B. 1978. The eyes of mesopelagic crustaceans. II. Streetsia challengeri (Amphipoda). Cell and Tissue Research 186:337–349. Meyer-Rochow, V.B., and S. Walsh. 1978. The eyes of mesopelagic crustaceans. III. Thysanopoda tricuspidata (Euphausiacea). Cell and Tissue Research 195:59–79. Nalbach, H.-O. 1990. Multisensory control of eyestalk orientation in decapod crustaceans: an ecological approach. Journal of Crustacean Biology 10:382–399. Nilsson, D.-E. 1982. The transparent compound eye of Hyperia (Crustacea): examination with a new method for analysis of refractive index gradients. Journal of Comparative Physiology A 147:339–349. Nilsson, D.-E. 1983. Evolutionary links between apposition and superposition optics in crustacean eyes. Nature 302:818–821. Nilsson, D.-E. 1989. Optics and evolution of the compound eye. Pages 30–73 in R.C. Hardie and D.G. Stavenga, editors. Facets of vision. Springer, Berlin Heidelberg. Nilsson, D.-E. 1995. Eye design, vision and invisibility in planktonic invertebrates. Pages 149–162 in P. Lenz, D. Hartline, J. Purcell, and D. Macmillan, editors. Zooplankton: sensory ecology and physiology. Gordon and Breach, Amsterdam. Nilsson, D.-E., and R.F. Modlin. 1994. A mysid shrimp carrying a pair of binoculars. Journal of Experimental Biology 189:213–236. Nilsson, D.-E., and H.L. Nilsson. 1981. A crustacean compound eye adapted for low light intensities (Isopoda). Journal of Comparative Physiology A 68:105–110. Nilsson, D.-E., E. Hallberg, and R. Elofsson. 1986. The ontogenetic development of refracting superposition eyes in crustaceans: transformation of optical design. Tissue and Cell 18:509–519. O’Neill, P.J., R.N. Jinks, E.D. Herzog, B.A. Battelle, L. Kass, G.H. Renninger, and S.C. Chamberlain. 1995. The morphology of the dorsal eye of the hydrothermal vent shrimp, Rimicaris exoculata. Visual Neuroscience 12:861–875. Pechenik, J.A. 1999. On the advantages and disadvantages of larval stages in benthic marine invertebrate life cycles. Marine Ecology Progress Series 177:269–298. Pelli, D.G., and S.C. Chamberlain. 1989. The visibility of 350 °C black-body radiation by the shrimp Rimicaris exoculata and man. Nature 337:460–461. Porter, M.L., M.J. Bok, P.R. Robinson, and T.W. Cronin. 2009. Molecular diversity of visual pigments in Stomatopoda (Crustacea). Visual Neuroscience 26:255–266. Porter, M.L., Y. Zhang, S. Desai, R.L. Caldwell, and T.W. Cronin. 2010. Evolution of anatomical and physiological specialization in the compound eyes of stomatopod crustaceans. Journal of Experimental Biology 213:3473–3496. Putkunz, C.T., and A.G. Peele. 2009. Detailed simulation of a Lobster-eye telescope. Optics Express 17:14156–14165.

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Thomas W. Cronin and Kathryn D. Feller Rajkumar, P., S.M. Rollman, T.A. Cook, and J.E. Layne. 2010. Molecular evidence for color discrimination in the Atlantic sand fiddler crab, Uca pugilator. Journal of Experimental Biology 213:4240–4248. Roberts, N.W., T.-H. Chiou, N.J. Marshall, and T.W. Cronin. 2009. A biological quarter-wave retarder with excellent achromaticity in the visible wavelength region. Nature Photonics 3:641–644. Sakamoto, K., O. Hisatomi, F. Tokunaga, and E. Eguchi. 1996. Two opsins from the compound eye of the crab Hemigrapsus sanguineus. Journal of Experimental Biology 199:441–450. Smith, F.E., and E.R. Baylor. 1953. Color responses in the Cladocera and their ecological significance. American Naturalist 87:49–55. Smith, K.C., and E.R. Macagno. 1990. UV photoreceptors in the compound eye of Daphnia magna (Crustacea, Branchiopoda). A fourth spectral class in single ommatidia. Journal of Comparative Physiology A 166:597–606. Smolka, J., and J.M. Hemmi. 2009. Topography of vision and behaviour. Journal of Experimental Biology 212:3522–3532. Stearns, D.E., and R.B. Forward Jr. 1984. Photosensitivity of the marine copepod, Acartia tonsa. Marine Biology 82:85–89. Van der Velden, J., Y. Zheng, B.W. Patullo, and D.L. Macmillan. 2008. Crayfish recognize the faces of fight opponents. PLoS ONE 3(2):e1695. doi: 10.1371/journal.pone.0001695 Van Dover, C.L., E.Z. Szuts, S.C. Chamberlain, and J.R. Cann. 1989. A novel eye in “eyeless” shrimp from hydrothermal vents of the Mid-Atlantic Ridge. Nature 337:458–460. Warrant, E., K. Bartsch, and C. Günther. 1999. Physiological optics in the hummingbird hawkmoth: a compound eye without ommatidia. Journal of Experimental Biology 202:497–511. Waterman, T.H. 1981. Polarization sensitivity. Pages 281–469 in H. Autrum, editor. Comparative physiology and evolution of invertebrates, B, Invertebrate visual centers and behavior, I. Handbook of sensory physiology VII/6B. Springer, New York. Zeil, J., and J.M. Hemmi. 2006. The visual ecology of fiddler crabs. Journal of Comparative Physiology A 192:1–25

10 THE CHEMICAL SENSES AND CHEMOSENSORY ECOLOGY OF CRUSTACEANS

Charles D. Derby and Marc J. Weissburg

Abstract The chemical world of crustaceans is complex and rich in information about features of the environment in which they are found. Consequently, it is not surprising that the chemical senses of crustaceans are used in most aspects of their lives, including reproduction, securing resources, interacting with conspecifics, and avoiding predators. The chemical senses serve some fundamental functions across species: they allow discrimination of the many biologically relevant molecules to help crustaceans determine the nature of the objects releasing those chemicals, and they determine spatiotemporal features of chemical plumes so the crustaceans can locate these objects in their environments. The chemical senses of crustaceans also show specialized adaptations that can vary with phylogeny, lifestyle, and habitat. Thus, there is a diversity of chemoreceptors in crustaceans, both within a given species and across species. Some homologous structures are known, such as aesthetasc sensilla, but the nature of the diversity of forms is often currently unknown. The chemical senses of crustaceans are organized into multiple pathways that differ in organization and function; these include olfaction and distributed chemoreception. Neuroscientists have used a few crustacean species, especially from the decapods, to study the form and function of chemosensors. Chemoreceptor neurons have been studied to understand mechanisms of sensory transduction and coding. Interneurons and circuits have been studied to understand central processing. In recent years, we have learned much about crustacean chemoreception, and new technologies promise to help us understand much more about the nature and function of their chemical senses.

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THE CHEMICAL WORLD OF CRUSTACEANS The chemical senses are rapidly adapting and evolving senses. Most stimuli, including visual, electrical, geomagnetic, and mechanical, are quantifiable physical energies, defined by wavelength and other physical properties that vary on a continuous scale. Some wavelengths may dominate in specific environments (e.g., red colors dominate in the deep sea because shorter wavelengths are filtered and longer wavelengths including red light penetrate deeper), but there are no new wavelengths under different conditions. Chemicals, on the other hand, can be newly created by organisms (e.g., as metabolic products) and can be used in diverse combinations to form complex mixtures of almost limitless variety. Thus, chemicals and the molecules involved in the detection of them, such as receptor genes, may be susceptible to more rapid evolutionary changes than most other stimuli and senses (Bargmann 2006, Symonds and Elgar 2008). Chemicals can be signals or cues. Signals are stimuli used in communication, in which emission and response have evolved for specific effects and for which the response is adaptive to the sender. Signals include pheromones such as those used in sexual contexts. Cues, on the other hand, are chemicals that also can affect many behaviors, but their emission has not evolved for the benefit of the sender but rather determined by selective pressure on the receiver only. Blood from injured animals may attract a predatory or scavenging crustacean, but the chemicals in them have not evolved for that function, and thus this stimulus is a cue rather than a signal. Wyatt (2010) proposes the term “signature mixtures” for chemicals released from an organism and whose response to it must be learned, as opposed to the innate responses to pheromones. Signature mixtures may include chemical mixtures used by members of a social species to differentiate among each other. Chemicals are used as stimuli for many functions and in diverse environments. Chemicals are associated with most biologically important, life-maintaining and -perpetuating processes, and consequently, animals, including crustaceans, use chemicals as integral parts of their lives. Pheromones are used in many groups of crustaceans and for many functions (see contributions in Breithaupt and Thiel 2011). Pheromones and/or signature mixtures can aid in recognizing species, sex, genetic relatedness, individual identity, social status, or aggressiveness. Pheromones may function either from a distance or on physical contact (Bauer 2011, Snell 2011). Despite the ubiquity of crustacean pheromones, their chemical identity remains elusive. For example, the chemical identity of sex pheromones largely remains unknown (Hardege and Terschak 2011, Kamio and Derby 2011) even for commercially important species for which there are obvious economic advantages from identifying the sex pheromones. Identification of water-borne sex pheromones has been particularly challenging, but some contact sex pheromones have been identified, such as α 2-macroglobulins in the copepod Tigriopus japonicus (Snell 2011). Cues are used in a wide assortment of functions and behaviors, including settlement (Clare 2011), camouflage (Stachowicz and Hay 1999), finding home or shelter (Rittschof and Cohen 2004, Briones-Fourzán et al. 2008, Lecchini et al. 2010), finding symbiotic hosts (Ambrosio and Brooks 2011), hatching of larvae (Rittschof and Cohen 2004), avoiding predators (Hay 2009, Hazlett 2011), identifying injured or diseased conspecifics (Behringer et al. 2008, Hazlett 2011), finding and consuming food (Derby and Zimmer 2012), and many more. The molecular identities of many cues have been elucidated, and here we mention only a few. One example is the cues mediating larval settlement. Larval barnacles must choose a settlement site near conspecifics because adult barnacles are immobile and have internal fertilization. Larval barnacles use multimodal cues in making their choice, with contact chemical cues from conspecifics being important. Building on a long history of research, Clare and colleagues identified the chemotactile cue to be α 2-macroglobulins, which are cuticular proteins in the integument of adult barnacles (Clare 2011). A second example is cues involved in selection of food by crustaceans. These



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include both feeding inducers and feeding deterrents. Feeding inducers are typically mixtures rather than single compounds, which is expected given that single chemicals are rarely if ever encountered in nature, and mixtures generally have higher information content. Crustaceans can analyze and discriminate among mixtures based on their components, even for mixtures that are closely related and vary only in the ratio of their components (Derby 2000). What is the basis for these behavioral responses to food? What are the bioactive compounds? Differences in sensitivity across crustacean species are partly attributable to the habitat in which they live. For aquatic crustaceans, feeding cues typically are water-soluble compounds, but lipophilic molecules can be relevant and are sometimes used as cues, particularly at close range (e.g., defensive compounds). For terrestrial crustaceans, both volatiles and water soluble molecules are relevant: the antennules are sensitive to volatile chemicals, and the legs and mouthparts are sensitive to water-soluble stimuli (Rittschof and Sutherland 1986). Differences in trophic levels also account for some of the interspecific differences in sensitivity to feeding cues. For example, carnivores are sensitive to nitrogen-containing compounds, such as amino acids, amines, nucleotides, and peptides, which are prevalent in tissues of their prey, and they are relatively insensitive to carbohydrates and sugars (Carr 1988, Zimmer-Faust 1993). Herbivores and omnivores, such as fiddler crabs (Weissburg and Zimmer-Faust 1991, Weissburg 1999), ghost crabs (Trott and Robertson 1984), kelp crabs (Zimmer-Faust and Case 1982), and crayfish (Tierney and Atema 1988, Corotto and O’Brien 2002), tend to be sensitive to sugars found in plants, bacteria, and diatoms, as well as to some amino acids. Specialist consumers often use specific molecules to decide which foods to eat. For example, the crab Caphyra rotundifrons, which feeds exclusively on the green alga Chlorodesmis fastigiata, is stimulated to feed by the compound chlorodesmin found in this seaweed, which also serves as a feeding deterrent against herbivorous fish (Hay et al. 1989). This adaptation for specialists to hone in on compounds that plants or algae use in defense is common in crustaceans, insects, and other arthropods (Eisner et al. 2007, Hay 2009). Feeding by generalist herbivorous and omnivorous crustaceans also is regulated by plant secondary metabolites that function as defensive compounds (Prusak et al. 2005, Parker et al. 2007, Hay 2009, ­chapter 18 in volume 2). Despite numerous studies concluding that dissolved free amino acids (DFAAs) play a primary role as attractants, this hypothesis is in revision (Derby and Zimmer 2012). Certainly, DFAAs display qualities consistent with an attractive role: they are highly soluble in seawater, stimulate sensory neurons, and evoke oriented locomotion. However, DFAAs released at rates measured from live and intact prey did not attract predators from a distance (Zimmer et al. 1999, Finelli et al. 2000). Although chemicals are released at much higher rates from recently injured prey or fresh carrion, compared with live and intact prey, field sites baited with DFAAs released at these ecologically relevant rates also did not attract predatory crustaceans from distances greater than 30 cm (Zimmer-Faust and Case 1982, Finelli et al. 2000). They did, however, significantly attract from those distances scavenging crustaceans such as the isopod Cirolana diminuta (Zimmer et  al. 1999). At very close range ( 10 cm/sec), and they occupy habitats with different fluid properties. Many aspects of the morphology and physiology of crustacean chemical sensing may only be interpretable in light of the fluid physical environment and consequent chemical milieu. Characterizing the structure of environmental chemicals requires information about spatial and temporal patterns of those chemicals. However, basic fluid dynamical patterns provide general guides to the forces that distribute chemicals and the corresponding information available to the animals. Recent reviews cover the hydrodynamics of chemical signaling in detail (Webster and Weissburg 2009, Weissburg 2011), but general aspects of flow and chemical environment are revealed by the Reynolds number, Re. This parameter describes the relative importance of inertial vs. viscous forces in fluids. Low Re values indicate laminar flow where parcels of fluid move in the direction of bulk flow and flow streamlines do not cross. A high Re indicates turbulent flow where there are unpredictable variations in flow velocity and direction. Diffusion is important in low Re environments, which produce predictable concentration gradients. High Re values denote that high flows and turbulent transport produce chaotic and unpredictable signals. Planktonic forms, such as copepods, anostracans, cladocerans, and mysids, typically live in low (1–10) Re regimes (Yen 2013). The defining features of stimuli in such environments are spatial coherence and strong gradients in stimulus intensity (Fig. 10.6B,C). Pheromones in the wake of moving female copepods or plumes in the wake of sinking particles are well-defined trails where biologically meaningful concentrations may persist tens of seconds after the chemical source has passed (Bagøien and Kiørboe 2005, Yen and Lasley 2011). Chemical gradients are particularly sharp in the axial direction, that is, the direction perpendicular to the long axis of the trail or plume. Feeding currents produced by copepods draw chemicals from around potential



The Chemical Senses and Chemosensory Ecology of Crustaceans

food into elongated “clouds” that pass over chemosensory sensilla on antennules and mouthparts (Fig. 10.6B). Benthic copepods exist in low Re environments due to their small size, but also because fluid velocity is reduced near a fluid interface, that is, within the boundary layer (Vogel 1994). Chemicals around such animals likely exist as a diffusive cloud, with a fairly sharp gradient. Larger crustaceans such as decapods experience Re values from 103 to 105, indicating that impacts of diffusion are relatively weak. Chemical distributions in these environments are probably the best characterized among those relevant to crustaceans. Turbulent plumes exhibit substantial and unpredictable spatial and temporal variations as a result of the chaotic nature of the flow, with filaments of high concentration interspersed with unscented fluid (Fig. 10.6A). The filamentous structure is particularly strong near the source, whereas the plume eventually becomes more homogeneous and dilute as it expands downstream. The range of Re values for semiterrestrial crustaceans is similar to their aquatic relatives; although the kinematic viscosity is lower, flow rates are higher, and the turbulent nature of airborne plumes is documented by studies on terrestrial insects (Murlis et al. 1992). In aquatic realms at least, turbulent plumes also exhibit substantial vertical variation. Regions closest to the bed (e.g., within the first several mm) are well-mixed, homogeneous, and diluted relative to elevated regions, where the stimulus structure displays greater variability (Jackson et al. 2007). Thus, sensors on different body regions of macroscopic, benthic crustaceans are challenged with different stimulus patterns. Some crustaceans use chemicals during social interactions, such as when rivals meet to establish dominance hierarchies (Berry and Breithaupt 2010). Animals in dominance contests produce chemical stimuli by ejecting urine from small pores at relatively high velocity to form a turbulent jet. These are well-mixed and homogeneous structures. Bivalves also produce jets of released metabolites, which can be used by crustaceans to locate prey (Smee and Weissburg 2006). However, the relatively low release velocity of these jets means that their characteristics are influenced mostly by turbulent processes occurring in ambient flow. Purely diffusive transport without flow is important only when transport distances are extremely small, which occurs mainly around chemosensory sensilla, that is, in the final phase of transport. Chemicals trapped within aesthetasc tufts retain the spatial properties imposed on the chemical plume by turbulent processes even as the chemicals reach the aesthetascs’ surface via diffusion (Koehl 2011). Critical aspects of spatial and temporal features of chemicals remain unknown, despite the fluid dynamical framework. Plumes with low Re values created by moving plankton have not been studied. Here, the low-flow environment may provide opportunities for modeling the structure of chemical plumes. Interaction of neighboring sources of chemicals creates complex patterns that remain unexamined. This may be consequential when neighboring sources of chemicals have different salience for the animals, such as attractive and aversive cues. Experiments with terrestrial arthropods suggest that the spacing and release characteristics of coupled attractive and aversive stimuli have large impacts on search behavior (Fadamiro et al. 1999). Lastly, we have concentrated on water-borne chemicals since crustaceans receive much information in this way. However, most crustaceans have chemosensors on the tips of their legs and claws that may be important as they approach the source. Sediment-associated chemicals are undoubtedly important in the lives of semiterrestrial crustaceans such as the gecarcinids and ocypodids. The distribution and characteristics of chemicals associated with sediments remains unknown despite their potential importance to food location. Crustacean chemosensors, like other sensors, respond to constant stimulation with decreases in firing rate over a number of different timescales, that is, neural adaptation of sensory responses (see examples of neural adaptation in ­chapters 3 and 8 in this volume). Neural

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Fig. 10.6. Structure of chemical signals experienced by large and small crustaceans. (A) Chemical plume created by releasing dye from a nozzle located 2.5 cm above the bed in a 5 cm/sec flow. The roughness Re of this flow is approximately 3, indicating smooth turbulent flow. Field of view is approximately 80 × 20 cm (l × w). (B) Chemicals drawn into the feeding current of the copepod Pleuromamma xiphias, measured using a chemical microelectrode. The rostrum of the copepod is at the lower right-hand corner, and the chemical was released from a pipette tip denoted by the circle on the upper left. Counter lines represent 10% decrements of the maximal center concentration (from Moore et al. 1999, with permission from the American Society of Limnology and Oceanography). (C)  Model of chemical concentration wake of a swimming plankton. The lowest concentration of this wake is defined as 10% of the maximal concentration. From Bearon and Magar 2010, with permission from Oxford University Press.

adaptation of sensory cells occurs in response to either constant stimulation (from hundreds of milliseconds to several seconds) or iterative pulses of stimuli (from milliseconds to seconds) presented at inter-pulse intervals from milliseconds to several seconds. Neural adaptation is different from habituation, which is also expressed as a decreasing response to repeated stimulation. However, habituation is not due to adaptation of sensory responses, as shown by the fact that the habituated response can be removed, or dishabituated, by presenting a stimulus from another sensory modality (see examples of habituation in ­chapters 2, 8, 11, 15, 18, and 19 in this volume). Studies on temporal dynamics in crustacean chemosensory neurons often are motivated by revealing relationships between physiological processes of adaptation-disadaptation and the ability of crustaceans to follow chemical plumes. These investigations revealed general features of neurons that seem useful in encoding temporal properties of stimuli. However, a strong mechanistic understanding of the role of dynamic neuronal properties in shaping responses to time-varying stimuli typically experienced by crustaceans is needed. One challenge is to use realistic temporal stimulus profiles when investigating neural responses. This is done rarely and only with certain chemical properties (e.g., Gomez et al. 1994). Most protocols use arbitrary pulse lengths and interstimulus intervals, with the former being very long compared to naturally occurring signals (Webster and Weissburg 2001). A second challenge is a lack of parallelism between physiological studies and the importance of sensory neurons in navigation. In spiny



The Chemical Senses and Chemosensory Ecology of Crustaceans

lobsters, which provide the bulk of physiological data, there is still considerable uncertainty as to which sensors on the antennules mediate navigation, though it is known that both aesthetascs and distributed chemoreceptors on the antennules can drive orientation (Caprio and Derby 2008, Schmidt and Mellon 2011). Atema and colleagues examined adaptation of glutamate-sensitive chemoreceptor neurons on legs of H. americanus, using pulses of various durations. Neurons showed reduced firing rate within the first 100–300 msec of long pulses (one to several sec) such that firing rate declines to background levels (Borroni and Atema 1988). Neurons challenged with these stimuli showed concentration-response functions that were right-shifted relative to their previous state. Similar effects were produced when these neurons were challenged with 300-msec pulses at intervals ranging from 5 to 20 sec (Voigt and Atema 1990). One consequence of this recalibration is to keep the dynamic range of neurons within the range of ambient signals. In addition, firing rates are most responsive to the ratio between background and pulse concentrations as opposed to absolute concentrations, strongly suggesting that neurons are encoding contrast as opposed to absolute intensity. Shifting concentration-response profiles and contrast encoding are helpful when determining the information in turbulent chemical plumes, since the concentration of chemical pulses increases toward the source (Webster and Weissburg 2001). In addition, turbulent chemical plumes are highly variable, and different animals tracking through them experience vastly different chemical concentrations (Page et al. 2011). This high degree of variability makes the encoding of absolute concentrations considerably less useful and requires either large working ranges or the ability to shift the working range as needed. Rates of adaptation-disadaptation, as well as other dynamic processes such as second messenger transduction cascades, set limits on the ability of neurons to respond to time-varying chemical stimuli. Antennular chemosensory neurons may require 200 msec to accurately encode stimulus intensity (Gomez and Atema 1996a). Neurons are capable of phase-locking to 100-msec pulses delivered at rates up to 1 Hz, and some neurons can phase-lock at 4 Hz, particularly with less concentrated stimuli (Gomez et al. 1994) (Fig. 10.7). The overall population response indicates flicker-fusion at frequencies above 2 Hz. Slow recovery rates (Gomez and Atema 1996b) reinforce the speculation that these chemosensors are designed to detect rapid and transient signals. The temporal dynamics of responses to pulses of chemicals seem to reflect the temporal structure of chemical plumes produced by prey or other objects. These neuronal processes also might constrain the information encoded by the periphery and consequently affect how decisions are made during navigation and orientation. Pulse frequencies in typical turbulent chemical plumes are about 1 Hz (Mead et al. 2003, Page et al. 2011) and less than 200 msec in duration (Webster and Weissburg 2001). Thus, physiological frequencies seem to be matched to the environment. The apparent difficulty in extracting much information from brief pulses is ameliorated by how crustaceans use such information. Blue crabs, at least, use a binary encoding scheme where suprathreshold filaments of chemicals are used to provide information in turbulent plumes, with thresholds scaled to the prevailing stimulus levels (Page et al. 2011). The lack of graded responses to chemical concentration reduces the need for the periphery to encode stimulus intensity precisely. Little comparative information exists on the dynamics of adaptation-disadaptation across animals, sensors, or with reference to other processes (e.g., encoding chemical quality). Chemosensory neurons in legs and claws of fiddler crabs can maintain firing rates even when exposed to 10-sec pulses of strong (10 mM) stimuli (Weissburg 1999). This is presumably adaptive for animals that are continuously exposed to sediment-bound chemical cues, which may be concentrated in comparison to average background concentrations in sea water. Chemosensory

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neurons on maxillipeds of spiny lobsters display properties intermediate between those on legs of fiddler crabs and antennules of lobsters, showing strong decreases in firing rate over 0.5 to 3 sec (Garm et al. 2005). The significance of this time course, as well as the observed species specificity, remains unresolved.

FUTURE DIRECTIONS There are many exciting areas for future study of crustacean chemoreception. Some are ripe for analysis due to technical advances that provide opportunities heretofore unavailable. We offer a few examples. 1. Behavior of crustaceans can be complex, sophisticated, and intricate. Equipment for digitally capturing and analyzing images has made it easier to study behaviors of all types including very fast behaviors, behaviors performed in the dark, and behavior in the field. 2. Chemical, physical, and biological variables are rarely static in nature. To be sure that their experiments are ecologically relevant, researchers should use appropriate “dynamic scaling” by stimulating with chemical and physical cues whose features realistically represent the changing native signaling environment of the animals of interest. Field instrumentation for monitoring flow habitats, and the increasing availability of flow tanks and flow visualization techniques, allow investigators to incorporate realistic environments and signals. 3. A challenge for studying chemical senses is to identify bioactive molecules. Sophisticated, reliable, and rapid techniques to separate and identify the molecular structures even in



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The Chemical Senses and Chemosensory Ecology of Crustaceans minute amounts are now available. This permits identification of bioactive molecules even when they are present at nanomolar concentrations and embedded in complex mixtures. With candidate chemical cues or signals in hand, use of reliable, high-throughput bioassays such as calcium imaging of activity of many cells simultaneously and fast behavioral assays will facilitate screening for activity. Ultimately, candidate cues or signals need to be tested in behavioral assays, preferably under as natural contexts as possible, even in the field. The genome of Daphnia, the first crustacean genome to be sequenced, was released in 2010, and now more crustacean species need to be sequenced, especially models for chemoreception research. Sequencing of targeted organs to generate transcriptomes for chemosensory organs, brains, and key species will also be important. Together with techniques to change expression levels, such as RNAi or knock-outs, functions of genes and their products can be defined. Optogenetics—the targeting of fast, light-activated channels, such as channelrhodopsin for activation and halorhodopsin for inactivation, for expression in cells containing molecules of interest—has revealed functions of cells and circuits in several species (Deisseroth 2011) and will do so when applied to crustaceans. Such genetic engineering can be used to activate specific types of neurons at will and might be used to activate chemosensory neurons expressing a particular receptor type, or other molecules, in behaving animals. The fields of sensory ecology and neuroecology combine techniques spanning from biophysical and molecular properties of nerve and muscle cells to community-wide impacts of trophic interactions, to undertand the consequences that neural mechanisms of sensory systems may have on ecological interactions (Dusenbery 1992, Zimmer and Derby 2011). These fields can help understand principles controlling the organization of communities by examining the processes of information transfer, the ability of animals to respond to signals indicating the presence of competitors, mates, food, dwelling sites, and other resources, and the effect of subsequent responses on community members.

SUMMARY The chemical senses of crustaceans play a role, often critical, in many of the fundamental problems facing them, including finding resources such as shelter, food, and mates, avoiding dangers such as toxins, predators, competitors, and aggressive or diseased conspecifics. Given their vast diversity, crustaceans are excellent organisms to use in comparative studies to reveal evolutionary adaptations to diverse habitats and lifestyles. Some species of crustaceans also have served as important model organisms in the study of basic principles of organization and function of chemosensory nervous systems, including sensory transduction, neural control of behavior, organization of central pathways, and adult neurogenesis. They have also been used to elucidate mechanisms underlying chemosensory discrimination of chemical mixtures and locating objects in their environments.

ACKNOWLEDGMENTS We thank our colleagues who have contributed to work in this field. We also thank funding agencies for their support over the years, currently from National Science Foundation grant IOS-1036742, The Plum Foundation John E.  Dowling Fellowship Fund, and the Colwin Endowed Summer Research Fellowship Fund from the Marine Biological Laboratory.

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REFERENCES Ache, B.W. 2002. Crustaceans as animal models for olfactory research. Pages 189–199 in K. Wiese, editor. Crustacean experimental systems in neurobiology. Springer, Berlin. Aggio, J.F., R. Tieu, A. Wei, and C.D. Derby. 2012. Oesophageal chemoreceptors of blue crabs, Callinectes sapidus, sense chemical deterrents and can block ingestion of food. Journal of Experimental Biology 215:1700–1710. Altner, I., H. Hatt, and H. Altner. 1983. Structural properties of bimodal chemo- and mechanosensitive setae on the pereiopod chelae of the crayfish, Austropotamobius torrentium. Cell and Tissue Research 228:357–374. Ambrosio, L.J., and W.R. Brooks. 2011. Recognition and use of ascidian hosts, and mate acquisition by the symbiotic pea crab Tunicotheres moseri (Rathbun, 1918): the role of chemical, visual and tactile cues. Symbiosis 53:53–61. Atema, J., and M.A. Steinbach. 2007. Chemical communication and social behavior of the lobster, Homarus americanus, and other decapod Crustacea. Pages 115–144 in J.E. Duffy and M. Thiel, editors. Evolutionary ecology of social and sexual systems: crustaceans as model organisms. Oxford University Press, Oxford. Atema, J., and R. Voigt. 1995. Behavior and sensory biology. Pages 331–348 in J.R. Factor, editor. Biology of the lobster, Homarus americanus. Academic, San Diego. Bagøien, E., and T. Kiørboe. 2005. Blind dating—mate finding in planktonic copepods. I. Tracking the pheromone trail of Centropages typicus. Marine Ecology Progress Series 300:105–115. Bargmann, C.I. 2006. Comparative chemosensation from receptors to ecology. Nature 444:295–301. Bauer, R.T. 2011. Chemical communication in decapod shrimps: the influence of mating and social systems on the relative importance of olfactory and contact pheromones. Pages 277–296 in T. Breithaupt and M. Thiel, editors. Chemical communication in crustaceans. Springer, New York. Bauer, R.T., and J.L. Caskey. 2006. Flagellar setae of the second antennae in decapod shrimps: sexual dimorphism and possible role in detection of contact sex pheromones. Invertebrate Reproduction and Development 49:51–60. Bauer, U., and H. Hatt. 1980. Demonstration of three different types of chemosensitive units in the crayfish claw using a computerized evaluation. Neuroscience Letters 17:209–214. Bearon, R.N., and V. Magar. 2010. Simple models of the chemical field around swimming plankton. Journal of Plankton Research 32:1599–1608. Behringer, D.C., M.J. Butler, and J.D. Shields. 2008. Ecological and physiological effects of Pav1 infection on the Caribbean spiny lobster (Panulirus argus Latreille). Journal of Experimental Marine Biology and Ecology 359:26–33. Belanger, R., X. Ren, K. McDowell, S. Chang, P. Moore, and B. Zielinski. 2008. Sensory setae on the major chelae of male crayfish, Orconectes rusticus (Decapoda, Astacidae)—impact of reproductive state on function and distribution. Journal of Crustacean Biology 28:27–36. Beltz, B.S., K. Kordas, M.M. Lee, J.B. Long, J.L. Benton, and D.C. Sandeman. 2003. Ecological, evolutionary, and functional correlates of sensilla number and glomerular density in the olfactory system of decapod crustaceans. Journal of Comparative Neurology 455:260–269. Berry, F.C., and T. Breithaupt. 2010. To signal or not to signal? Chemical communication by urine-borne signals mirrors sexual conflict in crayfish. BMC Biology 8:25. Blaustein, D.N., R.B. Simmons, M.F. Burgess, C.D. Derby, M. Nishikawa, and K.S. Olson. 1993. Ultrastructural localization of 5’AMP odorant receptor sites on the dendrites of olfactory receptor neurons of the spiny lobster. Journal of Neuroscience 13:2821–2828. Bobkov, Y.V., and B.W. Ache. 2007a. Block by amiloride derivatives of odor-evoked discharge in lobster olfactory receptor neurons through action on a presumptive TRP channel. Chemical Senses 32:149–159. Bobkov, Y.V., and B.W. Ache. 2007b. Intrinsically bursting olfactory receptor neurons. Journal of Neurophysiology 97:1052–1057. Borroni, P.F., and J. Atema. 1988. Adaptation in chemoreceptor cells. I. Self-adapting backgrounds determine threshold and cause parallel shift of response function. Journal of Comparative Physiology A 164:67–74.



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Boxshall, G.A., and R. Huys. 1998. The ontogeny and phylogeny of copepod antennules. Proceedings of the Royal Society of London B—Biological Sciences 353:765–786. Breithaupt, T., and M. Thiel, editors. 2011. Chemical communication in crustaceans. Springer, New York. Briones-Fourzán, P., E. Ramírez-Zaldívar, and E. Lozano-Álvarez. 2008. Influence of conspecific and heterospecific aggregation cues and alarm odors on shelter choice by syntopic spiny lobsters. Biological Bulletin 215:182–190. Caprio, J., and C.D. Derby. 2008. Aquatic animal models in the study of chemoreception. Pages 97–134 in S. Firestein and G.K. Beauchamp, editors. The senses, Vol. 4, Olfaction and taste. Academic Press, San Diego. Carr, W.E.S. 1988. The molecular nature of chemical stimuli in the aquatic environment. Pages 3–28 in J. Atema, R.R. Fay, A.N. Popper, and W.N. Tavolga, editors. Sensory biology of aquatic animals. Springer-Verlag, New York. Carr, W.E.S., R.A. Gleeson, and H.G. Trapido-Rosenthal. 1990. The role of perireceptor events in chemosensory processes. Trends in Neurosciences 13:212–215. Cate, H.S., and C.D. Derby. 2001. Morphology and distribution of setae on the antennules of the Caribbean spiny lobster Panulirus argus reveal new types of bimodal chemo-mechanosensilla. Cell and Tissue Research 304:439–454. Cate, H.S., and C.D. Derby. 2002. Hooded sensilla homologues: structural variations of a widely distributed bimodal chemo-mechanosensillum. Journal of Comparative Neurology 444:345–357. Clare, A.S. 2011. Toward a characterization of the chemical cue to barnacle gregariousness. Pages 431–450 in T. Breithaupt and M. Thiel, editors. Chemical communication in crustaceans. Springer, New York. Corey, E.A., Y. Bobkov, K. Ukhanov, and B.W. Ache. 2013. Ionotropic crustacean olfactory receptors. PLoS ONE 8:e60551. Corotto, F.S., and M.R. O’Brien. 2002. Chemosensory stimuli for the walking legs of the crayfish Procambarus clarkii. Journal of Chemical Ecology 28:1117–1130. Corotto, F., R. Voigt, and J. Atema. 1992. Spectral tuning of chemoreceptor cells of the third maxilliped of the lobster, Homarus americanus. Biological Bulletin 183:456–462. Croset, V., R. Rytz, S.F. Cummins, A. Budd, D. Brawand, H. Kaessmann, T.J. Gibson, and R. Benton. 2010. Ancient protostome origin of chemosensory ionotropic glutamate receptors and the evolution of insect taste and olfaction. PLoS Genetics 6:e1001064. Dahl, E., H. Emanuelsson, and C. von Mecklenburg. 1970. Pheromone reception in the males of the amphipod Gammarus duebeni Lilljeborg. Oikos 21:42–47. Deisseroth, K. 2011. Optogenetics. Nature Methods 8:26–29. Derby, C.D. 1989. Physiology of sensory neurons in morphologically identified cuticular sensilla of crustaceans. Pages 27–47 in B.E. Felgenhauer, L. Watling, and A.B. Thistle, editors. Functional morphology of feeding and grooming in Crustacea. A.A. Balkema Press, Rotterdam. Derby, C.D. 2000. Learning from spiny lobsters about chemosensory coding of mixtures. Physiology and Behavior 69:203–209. Derby, C.D., and J. Atema. 1988. Chemoreceptor cells in aquatic invertebrates: peripheral filtering mechanisms in decapod crustaceans. Pages 365–385 in J. Atema, R.R. Fay, A.N. Popper, and W.N. Tavolga, editors. Sensory biology of aquatic animals. Springer, New York. Derby, C.D., and R.K. Zimmer. 2012 Neuroecology of predator-prey interactions. Pages 158–171 in C. Brönmark and L.-A. Hansson, editors. Chemical ecology in aquatic systems. Oxford University Press, London. Dusenbery, D.B. 1992. Sensory ecology: how organisms acquire and respond to information. W.H. Freeman, New York. Eisner, T., M. Eisner, and M. Siegler. 2007. Secret weapons: defenses of insects, spiders, scorpions, and other many-legged creatures. Belknap Press of Harvard University Press, Cambridge, MA. Eisthen, H.L. 2002. Why are olfactory systems of different animals so similar? Brain Behavior and Evolution 59:273–293. Elofsson, R., and R.R. Hessler. 1991. Sensory morphology in the antennae of the cephalocarid Hutchinsoniella macracantha. Journal of Crustacean Biology 11:345–355.

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Charles D. Derby and Marc J. Weissburg Fadamiro, H.Y., A.A. Cosse, and T.C. Baker. 1999. Fine-scale resolution of closely spaced pheromone and antagonist filaments by flying male Helicoverpa zea. Journal of Comparative Physiology A 185:131–141. Finelli, C.M., N.D. Pentcheff, R.K. Zimmer, and D.S. Wethey. 2000. Physical constraints on ecological processes: a field test of odor-mediated foraging. Ecology 81:784–797. Förster, C., and J.A. Baeza. 2001. Active brood care in the anomuran crab Petrolisthes violaceus (Decapoda: Anomura: Porcellanidae): grooming of brooded embryos by the fifth pereiopods. Journal of Crustacean Biology 21:606-615. Forward, R.B., R.A. Tankersley, and D. Rittschof. 2001. Cues for metamorphosis of brachyuran crabs: an overview. American Zoologist 41:1108–1122. Galizia, C.G., and W. Rössler. 2010. Parallel olfactory systems in insects: anatomy and function. Annual Review of Entomology 55:399–420. Garm, A.L., and L. Watling. 2012. The crustacean integument: setae, scales, and other ornamentation. Pages 167–198 in L. Watling and M. Thiel, editors. The natural history of the Crustacea, Vol. 1. Functional morphology and diversity. Oxford University Press, New York. Garm, A.L, S. Shabani, J.T. Høeg, and C.D. Derby. 2005. Chemosensory neurons in the mouthparts of the spiny lobsters Panulirus argus and Panulirus interruptus (Crustacea: Decapoda). Journal of Experimental Marine Biology and Ecology 314:175–186. Ghiradella, H., T. Case, and J. Cronshaw. 1968. Structure of aesthetascs in select marine and terrestrial decapods: chemoreceptor morphology and environment. American Zoologist 8:603–621. Gleeson, R.A. 1982. Morphological and behavioral identification of sensory structures mediating pheromone reception in the blue crab, Callinectes sapidus. Biological Bulletin 163:162–171. Gleeson, R.A., and B.W. Ache. 1985. Amino acid suppression of taurine-sensitive chemosensory neurons. Brain Research 335:99–107. Gleeson, R.A., M.G. Wheatly, and C.L. Reiber. 1997. Perireceptor mechanisms sustaining olfaction at low salinities: insight from the euryhaline blue crab Callinectes sapidus. Journal of Experimental Biology 200:445–456. Gomez, G., and J. Atema. 1996a. Temporal resolution in olfaction: stimulus integration time of lobster chemoreceptor cells. Journal of Experimental Biology 199:1771–1779. Gomez, G., and J. Atema. 1996b. Temporal resolution in olfaction. II. Time course of recovery from adaptation in lobster chemoreceptor cells. Journal of Neurophysiology 76:1340–1343. Gomez, G., R. Voigt, and J. Atema.1994. Frequency filter properties of lobster chemoreceptor cells determined with high-resolution stimulus measurement. Journal of Comparative Physiology A 174:803–811. Grasso, R.W., and J.A. Basil. 2002. How lobsters, crayfishes, and crabs locate sources of odor: current perspectives and future directions. Current Opinion in Neurobiology 12:721–727. Grünert, U., and B.W. Ache. 1988. Ultrastructure of the aesthetasc (olfactory) sensilla of the spiny lobster, Panulirus argus. Cell and Tissue Research 251:95–103. Hallberg, E., and M. Skog. 2011. Chemosensory sensilla in crustaceans. Pages 85–102 in T. Breithaupt and M. Thiel, editors. Chemical communication in crustaceans. Springer, New York. Hallberg, E., K.U.I. Johansson, and R. Elofsson. 1992. The aesthetasc concept: structural variations of putative olfactory receptor cell complexes in Crustacea. Microscopy Research and Technique 22:325–335. Hallberg, E., K.U.I. Johansson, and R. Wallén. 1997. Olfactory sensilla in crustaceans: morphology, sexual dimorphism, and distribution patterns. International Journal of Insect Morphology and Embryology 26:173–180. Hamilton, K.A., K.A. Lindberg, and J.F. Case. 1985. Structure of dactyl sensilla in the kelp crab Pugettia producta. Journal of Morphology 185:349–366. Hardege, J.D., and J.A. Terschak. 2011. Identification of crustacean sex pheromones. Pages 373–392 in T. Breithaupt and M. Thiel, editors. Chemical communication in crustaceans. Springer, New York. Harrison, P.J.H., H.S. Cate, P. Steullet, and C.D. Derby. 2001. Structural plasticity in the olfactory system of adult spiny lobsters: postembryonic development permits life-long growth, turnover, and regeneration. Marine and Freshwater Research 52:1357–1366.



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Hatt, H., and B.W. Ache. 1994. Cyclic nucleotide- and inositol phosphate-gated ion channels in lobster olfactory receptor neurons. Proceedings of the National Academy of Sciences of the United States of America 91:6264–6268. Hay, M.E. 2009. Marine chemical ecology: chemical signals and cues structure marine populations, communities, and ecosystems. Annual Review of Marine Sciences 1:193–212. Hay, M.E., J.R. Pawlik, J.E. Duffy, and W. Fenical. 1989. Seaweed-herbivore-predator interactions: host-plant specialization reduces predation on small herbivores. Oecologia 81:418–427. Hazlett, B.A. 2011. Chemical cues and reducing the risk of predation. Pages 355–370 in T. Breithaupt and M. Thiel, editors. Chemical communication in crustaceans. Springer, New York. Helluy, S.M., J.L. Benton, K.A. Langworthy, M.L. Ruchhoeft, and B.S. Beltz. 1996. Glomerular organization in the developing olfactory and accessory lobes of the American lobster: stabilization of numbers and increase in size after metamorphosis. Journal of Neurobiology 29:459–472. Houser, L.T., and C.E. Epifanio. 2009. Impacts of biochemical cues on horizontal swimming behavior of individual crab larvae. Marine and Freshwater Behaviour and Physiology 42:249–264. Jackson, J.L., D.R. Webster, S. Rahman, and M.J. Weissburg. 2007. Bed roughness effects on boundary-layer turbulence and consequences for odor tracking behavior of blue crabs (Callinectes sapidus). Limnology and Oceanography 52:1883–1897. Johansson, K.U.I., and E. Hallberg. 1992. Male-specific structures in the olfactory system of mysids (Mysidacea; Crustacea). Cell and Tissue Research 268:359–368. Johansson, K.U.I., L. Gefors, R. Wallén, and E. Hallberg.1996. Structure and distribution patterns of aesthetascs and male-specific sensilla in Lophogaster typicus (Mysidacea). Journal of Crustacean Biology 16:45–53. Johnson, B.R., R. Voigt, P.F. Borroni, and J. Atema. 1984. Response properties of lobster chemoreceptors: tuning of primary taste neurons in walking legs. Journal of Comparative Physiology A 155:593–604. Kamio, M., and C.D. Derby. 2011. Approaches to a molecular identification of sex pheromones in blue crabs. Pages 393–412 in T. Breithaupt and M. Thiel, editors. Chemical communication in crustaceans. Springer, New York. Keller, T.A., I. Powell, and M.J. Weissburg. 2003. Role of appendages in chemically mediated orientation of blue crabs. Marine Ecology Progress Series 261:217–231. Kennish, M.J. 1986. Ecology of estuaries, Vol. 1, Physical and chemical aspects. CRC, Miami. Koehl, M.A.R. 2011. Hydrodynamics of sniffing by crustaceans. Pages 85–102 in T. Breithaupt and M. Thiel, editors. Chemical communication in crustaceans. Springer, New York. Kurbjeweit, F., and C. Buchholz. 1991. Structure and suspected functions of antennular sensilla and pores of three Arctic copepods (Calanus glacialis, Metridia longa, Pareuchaeta norvegica). Meeresforschung 33:168–182. Lazarini, F., and P.M. Lledo. 2011. Is adult neurogenesis essential for olfaction? Trends in Neurosciences 34:20–30. Lecchini, D., S.C. Mills, C. Brié, R. Maurin, and B. Banaigs. 2010. Ecological determinants and sensory mechanisms in habitat selection of crustacean postlarvae. Behavioral Ecology 21:599–607. Mead, K.S. 2008. Do antennule and aesthetasc structure in the crayfish Orconectes virilis correlate with flow habitat? Integrative and Comparative Biology 48:823–833. Mead, K.S., M.B. Wiley, M.A.R. Koehl, and J.R. Koseff. 2003. Fine-scale patterns of odor encounter by the antennule of the mantis shrimp tracking turbulent plumes in wave-affected and unidirectional flow. Journal of Experimental Biology 206:181–193. Mellon, DeF. Jr. 2000. Convergence of multimodal sensory input onto higher-level neurons of the crayfish olfactory pathway. Journal of Neurophysiology 84:3043–3055. Mellon, DeF. Jr., and V. Alones. 1993. Cellular organization and growth-related plasticity of the crayfish olfactory midbrain. Microscopy Research and Technique 24:231–259. Michel, W.C., T.S. McClintock, and B.W. Ache. 1991. Inhibition of lobster olfactory receptor cells by an odor-activated potassium conductance. Journal of Neurophysiology 65:446–453. Moore, P.A., D.M. Fields, and J. Yen. 1999. Physical constraints of chemoreception in foraging copepods. Limnology and Oceanography 44:166–177.

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Charles D. Derby and Marc J. Weissburg Murlis, J., J. S. Elkinton, and R. T. Cardé. 1992. Odor plumes and how insects use them. Annual Review of Entomology 37:505–532. Nakagawa, T., and L.B. Vosshall. 2009. Controversy and consensus: noncanonical signaling mechanisms in the insect olfactory system. Current Opinion in Neurobiology 19:284–292. Ohtsuka, S., and R. Huys. 2001. Sexual dimorphism in calanoid copepods: morphology and function. Hydrobiologia 453/454:441–466. Olsen, S.R., and R.I. Wilson. 2008. Lateral presynaptic inhibition mediates gain control in an olfactory circuit. Nature 452:956–960. Olson, K.S., and C.D. Derby. 1995. Inhibition of taurine 5’AMP olfactory receptor sites of the spiny lobster Panulirus argus by odorant compounds and mixtures. Journal of Comparative Physiology A 176:527–540. Page, J.L., D.B. Dickman, D.R. Webster, and M.J. Weissburg. 2011. Getting ahead: context-dependent responses to odorant filaments drives along-stream progress during odor tracking in blue crabs. Journal of Experimental Biology 214:1498–1512. Parker, J.D., D.E. Burkepile, D.O. Collins, J. Kubanek, and M.E. Hay. 2007. Mosses as chemically-defended refugia for freshwater macroinvertebrates. Oikos 116:302–312. Peñalva-Arana, D.C., M. Lynch, and H.M. Robertson. 2009. The chemoreceptor genes of the waterflea Daphnia pulex: many Grs but no Ors. BMC Evolutionary Biology 9:79. Prusak, A.C., J. O’Neal, and J. Kubanek. 2005. Prevalence of chemical defenses in freshwater macrophytes. Journal of Chemical Ecology 31:1145–1160. Rittschof, D., and J.H. Cohen. 2004. Crustacean peptide and peptide-like pheromones and kairomones. Peptides 25:1503–1516. Rittschof, D., and J.P. Sutherland. 1986. Field studies on chemically mediated behavior in land hermit crabs: volatile and nonvolatile odors. Journal of Chemical Ecology 12:1273–1284. Sandeman, D., R. Sandeman, C. Derby, and M. Schmidt. 1992. Morphology of the brain of crayfish, crabs, and spiny lobsters: a common nomenclature for homologous structures. Biological Bulletin 183:304–326. Sandeman, D.C., G. Scholtz, and R.E. Sandeman. 1993. Brain evolution in decapod Crustacea. Journal of Experimental Zoology 265:112–133. Sandeman, D.C., F. Bazin, and B.S. Beltz. 2011. Adult neurogenesis: examples from the decapod crustaceans and comparisons with mammals. Arthropod Structure and Development 40:258–275. Sandeman, R.E., and D.C. Sandeman. 1996. Pre-embryonic and postembryonic development, growth and turnover of olfactory receptor neurons in crayfish antennules. Journal of Experimental Biology 199:2409–2418. Sandoz, J.-C., N. Deisig, M.G. de Brito Sanchez, and M. Giurfa. 2007. Understanding the logics of pheromone processing in the honeybee brain: from labeled-lines to across-fiber patterns. Frontiers in Behavioral Neuroscience 1:5. Schachtner, J., M. Schmidt, and U. Homberg. 2005. Organization and evolutionary trends of primary olfactory brain centers in Tetraconata (Crustacea+Hexapoda). Arthropod Structure and Development 34:257–299. Schmidt, M. 1989. The hair-peg organs of the shore crab, Carcinus maenas (Crustacea, Decapoda): ultrastructural and functional properties of sensilla sensitive to changes in seawater concentration. Cell and Tissue Research 257:609–621. Schmidt, M. 2007. The olfactory pathway of decapod crustaceans—an invertebrate model for life-long neurogenesis. Chemical Senses 32:365–384 Schmidt, M., and C.D. Derby. 2005. Non-olfactory chemoreceptors in asymmetric setae activate antennular grooming behavior in the Caribbean spiny lobster, Panulirus argus. Journal of Experimental Biology 208:233–248. Schmidt, M., and W. Gnatzy. 1984. Are the funnel-canal organs the “campaniform sensilla” of the shore crab, Carcinus maenas (Decapoda, Crustacea)? II. Ultrastructure. Cell and Tissue Research 237:81–93. Schmidt M., and W. Gnatzy. 1989. Specificity and response characteristics of gustatory sensilla (funnel-canal organs) on the dactyls of the shore crab, Carcinus maenas (Crustacea, Decapoda). Journal of Comparative Physiology A 166:227–242.



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Schmidt, M., and DeF. Mellon Jr. 2011. Neuronal processing of chemical information in crustaceans. Pages 123–147 in T. Breithaupt and M. Thiel, editors. Chemical communication in crustaceans. Springer, New York. Schmitt, B.C., and B.W. Ache. 1979. Olfaction: responses of a decapod crustacean are enhanced by flicking. Science 205:204–206. Silbering, A.F., R. Okada, K. Ito, and C.G. Galizia. 2008. Olfactory information processing in the Drosophila antennal lobe: anything goes? Journal of Neuroscience 28:13075–13087. Simon, T.W., and C.D. Derby. 1995. Mixture suppression without inhibition for binary mixtures from whole cell patch clamp studies of in situ olfactory receptor neurons of the spiny lobster. Brain Research 678:213–224. Smee, D.L., and M.J. Weissburg. 2006. Clamming up: environmental forces diminish the perceptive ability of bivalve prey. Ecology 87:1587–1598. Snell, T. 2011. Contact chemoreception and its role in zooplankton mate recognition. Pages 451–466 in T. Breithaupt and M. Thiel, editors. Chemical communication in crustaceans. Springer, New York. Stachowicz, J.J., and M.E. Hay. 1999. Reducing predation through chemically-mediated camouflage: indirect effects of plant defenses on herbivores. Ecology 80:495–509. Stepanyan, R., K. Day, J. Urban, D.L. Hardin, R.S. Shetty, C.D. Derby, B.W. Ache, and T.S. McClintock. 2006. Gene expression and specificity in the mature zone of the lobster olfactory receptor organ. Physiological Genomics 25:224–233. Steullet, P., H.S. Cate, and C.D. Derby. 2000a. A spatiotemporal wave of turnover and functional maturation of olfactory receptor neurons in the spiny lobster Panulirus argus. Journal of Neuroscience 20:3282–3294. Steullet, P., H.S. Cate, W.C. Michel, and C.D. Derby. 2000b. Functional units of a compound nose: aesthetasc sensilla house similar populations of olfactory receptor neurons on the crustacean antennule. Journal of Comparative Neurology 418:270–280. Sullivan, J.M., and B.S. Beltz. 2004. Evolutionary changes in the olfactory projection neuron pathways of eumalacostracan crustaceans. Journal of Comparative Neurology 470:25–38. Sullivan, J.M., and B.S. Beltz. 2005. Integration and segregation of inputs to higher-order neuropils of the crayfish brain. Journal of Comparative Neurology 481:118–126. Symonds, M.R.E., and M.A. Elgar 2008. The evolution of pheromone diversity. Trends in Ecology and Evolution 23:220–228. Tadesse, T., M. Schmidt, W.W. Walthall, P.C. Tai, and C.D. Derby. 2011. Distribution and function of splash, an achaete-scute homolog in the adult olfactory organ of the Caribbean spiny lobster Panulirus argus. Developmental Neurobiology 71:316–335. Tierney, A.J., and J. Atema. 1988. Behavioral responses of crayfish (Orconectes virilis and Orconectes rusticus) to chemical feeding stimulants. Journal of Chemical Ecology 14:123–133. Tierney, A.J., C.S. Thompson, and D.W. Dunham. 1986. Fine-structure of aesthetasc chemoreceptors in the crayfish Orconectes propinquus. Canadian Journal of Zoology 64:392–399. Trott, T.J., and J.R. Robertson. 1984. Chemical stimulants of cheliped flexion behavior by the western Atlantic ghost crab Ocypode quadrata (Fabricius). Journal of Experimental Marine Biology and Ecology 78:237–252. Vieira, F.G., and J. Rozas. 2011. Comparative genomics of the odorant-binding and chemosensory protein gene families across the Arthropoda: origin and evolutionary history of the chemosensory system. Genome Biology and Evolution 3:476–490. Vogel, S. 1994. Life in moving fluids. Princeton University Press, Princeton, NJ. Voigt, R., and J. Atema. 1990. Adaptation in chemoreceptor cells. III. Effects of cumulative adaptation. Journal of Comparative Physiology A 166:865–874. Voigt, R., and J. Atema. 1992. Tuning of chemoreceptor cells of the second antenna of the American lobster (Homarus americanus) with a comparison of four of its other chemoreceptor organs. Journal of Comparative Physiology A 171:673–683. Wachowiak, M., and B.W. Ache. 1994. Morphology and physiology of multiglomerular olfactory projection neurons in the spiny lobster. Journal of Comparative Physiology A 175:35–48. Wachowiak, M., and B.W. Ache. 1998. Multiple inhibitory pathways shape odor-evoked responses in lobster olfactory projection neurons. Journal of Comparative Physiology A 182:425–434.

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292

Charles D. Derby and Marc J. Weissburg Wachowiak, M., L.B. Cohen, and B.W. Ache. 2002. Presynaptic inhibition of olfactory receptor neurons. Microscopy Research and Technique 58:365–375. Wachowiak, M., C.E. Diebel, and B.W. Ache. 1996. Functional organization of olfactory processing in the accessory lobe of the spiny lobster. Journal of Comparative Physiology A 178:211–226. Weatherby, T.M., K.K. Wong, and P.H. Lenz. 1994. Fine structure of the distal sensory setae on the first antennae of Pleuromamma xiphias Giesbrecht (Copepoda). Journal of Crustacean Biology 14:670–685. Webster, D.R., and M.J. Weissburg. 2001. Chemosensory guidance cues in a turbulent odor plume. Limnology and Oceanography 46:1048–1053. Webster, D.R., and M.J. Weissburg. 2009. The hydrodynamics of chemical cues among aquatic organisms. Annual Review of Fluid Mechanics 41:73–90. Weissburg, M.J. 1993. Sex and the single forager: gender-specific energy maximizing strategies in the fiddler crab Uca pugnax. Ecology 74:279–291. Weissburg, M.J. 1999. Sex-specific tuning and dose-response functions in male and female fiddler crabs, Uca pugnax. Journal of Comparative Physiology A 353:787–804. Weissburg, M.J. 2001. Sex, sensitivity, and second messengers: differential effect of cyclic nucleotide mediated inhibition in the chemosensory system of fiddler crabs. Journal of Comparative Physiology A 187:489–498. Weissburg, M.J. 2011. Waterborne chemical communication: stimulus dispersal dynamics and orientation strategies in crustaceans. Pages 63–83 in T. Breithaupt and M. Thiel, editors. Chemical communication in crustaceans. Springer, New York. Weissburg, M.J., and C.D. Derby. 1995. Regulation of sex-specific feeding behavior in fiddler crabs: physiological properties of chemoreceptor neurons in claws and legs of males and females. Journal of Comparative Physiology A 176:513–526. Weissburg, M.J., and R.K. Zimmer-Faust. 1991. Ontogeny versus phylogeny in determining patterns of chemoreception: initial studies with fiddler crabs. Biological Bulletin 181:205–215. Weissburg, M.J., J. Pearce, C.K. Govind, and C.D. Derby. 1996. Sexually dimorphic patterns of neural organization in the feeding appendages of fiddler crabs. Cell and Tissue Research 286:155–166. Weissburg, M.J., C.D. Derby, O. Johnson, B. McAlvin, and J.M. Moffett Jr. 2001. Transsexual limb transplants in fiddler crabs and the expression of novel sensory capabilities. Journal of Comparative Neurology 440:311–320. Wyatt, T.D. 2010. Pheromones and signature mixtures: defining species-wide signals and variable cues for individuality in both invertebrates and vertebrates. Journal of Comparative Physiology A 196:685–700. Yao, C.A., and J.R Carlson. 2010. Role of G-proteins in odor-sensing and CO2-sensing neurons in Drosophila. Journal of Neuroscience 30:4562–4572. Yen, J. 2013. Appendage diversity and modes of locomotion: swimming at intermediate Reynolds numbers. Pages 296–318 in L. Watling and M. Thiel, editors. The natural history of the Crustacea, Vol. 1. Functional morphology and diversity. Oxford University Press, New York. Yen, J., and R. Lasley. 2011. Chemical communication between copepods: finding the mate in a fluid environment. Pages 177–197 in T. Breithaupt and M. Thiel, editors. Chemical communication in crustaceans. Springer, New York. Zimmer, R.K., and C.D. Derby. 2011. Neuroecology and the need for broader synthesis. Integrative and Comparative Biology 51:751–755. Zimmer, R.K., J.E. Commins, and K.A. Browne. 1999. The regulatory effects of environmental chemical signals on search behavior and foraging success. Ecology 80:1432–1446. Zimmer-Faust, R.K., and J.F. Case. 1982. Organization of food search in the kelp crab, Pugettia producta (Randall). Journal of Experimental Marine Biology and Ecology 57:237–255. Zimmer-Faust, R.K. 1987. Crustacean chemical perception: towards a theory on optimal chemoreception. Biological Bulletin 172:10–29. Zimmer-Faust, R.K. 1993. ATP: a potent prey attractant evoking carnivory. Limnology and Oceanography 38:1271–1275.

11 MECHANORECEPTION IN CRUSTACEANS OF THE PELAGIC REALM

Petra H. Lenz and Daniel K. Hartline

Abstract Mechanoreception is used by pelagic crustaceans in the detection of predators, prey and mates, and communication. Behavioral studies have demonstrated how stimulus characteristics affect the response, that is, whether to attack, escape from, or pursue the source of the stimulus. However, the pelagic environment places constraints on the design of the receptors due to the physics of inhabiting a viscous environment with no fixed reference point. Thus, near-field disturbances are detected as differences in water flow between the sensor and the organism. These hydromechanic cues are transduced into a biological signal by cuticular sensilla, which in crustaceans are of scolopidial origin. The mechanosensory and supporting cells share many similarities, including distal and proximal anchoring of the dendrites separated by a stiff region. Modifications to the basic design are common, and these include a particularly stiff design in the copepods. Physiologically, the mechanoreceptors are characterized by high sensitivity, and they have a greater frequency range than is usual for sensilla of benthic decapods. The widespread occurrence of myelinated axons in both the Copepoda and some of the pelagic Malacostraca suggest that conduction velocities are important to shorten response times and improve stimulus localization.

INTRODUCTION Crustaceans and other organisms, whether of pelagic, benthic, or terrestrial habitat, use mechanoreceptors to obtain sensory information from the environment, as well as to provide internal 293

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Petra H. Lenz and Daniel K. Hartline information on appendage position and muscle stretch (Bush and Laverack 1982). Crustaceans are mostly aquatic or semi-aquatic organisms. They can be divided into two broad categories: pelagic and benthic. From the point of view of mechanoreception, this difference is a fundamental distinction with significant impact on neural mechanisms. Pelagic crustaceans inhabit a three-dimensional space, and detection of water movements is made without the fixed point of reference available to benthic organisms. Thus, the physics of near-field disturbances has a different impact on mechanoreception in pelagic crustaceans than on benthic forms. Behavioral studies of pelagic crustaceans have underscored the importance of mechanical cues in predator evasion, communication, and prey detection, while the differences in the physical and biological environment have led to specialized adaptations in mechanoreceptor physiology and morphology. This chapter focuses on mechanoreception in pelagic forms. Mechanoreceptors in crustaceans in general are described in ­chapter 3 of this volume, and for benthic crustaceans, mechanoreceptors involved in walking and posture are described in ­chapter 14 of this volume. Pelagic crustaceans are typically small, ranging between 1000 m; Fig. 11.1B), which are characterized by high pressure, no sunlight, constantly low temperatures (~4°C), and on average, scarce food resources, regardless of whether it is pelagic or benthic. In contrast, the better-studied surface-dwelling crustaceans inhabit the intertidal and subtidal benthic zones and shallow pelagic depths. Crustaceans, including the pelagic forms, possess an abundance of mechanoreceptors of several types. Reviews, primarily focused on benthic decapod forms, provide overviews of the types of mechanoreceptors present in the taxon, including structure-function considerations (Bush and Laverack 1982, ­chapter 3 in this volume). While much work has focused on proprioceptive mechanoreceptors, the chordotonal organs and muscle stretch receptors (Whitear 1962, Bush and Laverack 1982), there are almost no recent studies on these in pelagic crustaceans, nor much reason to expect pelagic-related specialization. Indeed, in the area of ecological interactions and sensory ecology, the cuticular hair or seta is the receptor type of greatest importance for these organisms. Thus the focus of this review will be on the form and function of setal exteroceptors and the role mechanoreception plays in the behavior and ecology of pelagic crustaceans.

PHYSICS OF PELAGIC MECHANORECEPTION Mechanoreception requires the relative physical movement of a receptor-activating structure with respect to a stretch-sensitive ion channel molecule. In nearly all cases, this is the movement of a detection structure such as a seta or otolith with respect to, or causing deformation of the receptor dendrite, which is populated with stretch-sensitive ion channels that underlie transduction (Goodman et al. 2004, Árnadóttir and Chalfie 2010). The coupling between this activating structure and the external medium through which the disturbance is transmitted imparts certain properties to the reception process, as well as providing limitations. There are four situations to consider: bulk flow (fluid flow on a scale larger than that of the organism), pressure cues



Mechanoreception in Crustaceans of the Pelagic Realm

Fig. 11.1. Pelagic crustaceans. See color version of this figure in centerfold. (A)  Euphausia sp., Malacostraca, length: 2–3 cm (photo © Steven Haddock by courtesy and with permission). (B) Gaetanus sp., juvenile, Copepoda, length:  3.5  mm (photo © Cheryl Clarke by courtesy and with permission. (C)  Acartia sp., adult female, Copepoda, length: 1.5 mm (photo © Ben Clauberg by courtesy and with permission, scale bar: 1 mm). (D) Spider crab megalopa, Malacostraca, length: 7 mm (photo courtesy of © Cheryl Clarke with permission). (E)  Bestiolina similis, nauplius stage 1, Copepoda, length:  70 μm (scanning electron micrograph, photo © Jennifer Kong by courtesy and with permission; scale bar: 50 μm).

(including far-field acoustic disturbances and hydrostatic pressure), near-field hydrodynamic disturbances (displacement of water at a source, spread according to hydrodynamic physics to neighboring parcels of water), and solid-object contact. As discussed below, bulk flow is difficult for pelagic organisms to detect, and direct evidence for far-field and hydrostatic detection in crustaceans in general is sparse, so consideration of these will be omitted (see ­chapter 3 in this volume). A good general reference for hydrodynamic disturbances is the review by Kalmijn (1988). Hydrodynamic Disturbances An organism suspended in a fluid is self-referenced in detecting hydrodynamic disturbances (fluid movement). Since the organism moves as a whole with the bulk flow in that environment,

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Petra H. Lenz and Daniel K. Hartline it cannot mechanically detect the relatively large disturbances associated with the movement without special adaptations. Detection of hydrodynamic cues in such environments depends on detection of relatively small water flows developed between the sensor and a reference part of the organism, often the hydrodynamic equivalent of the organism’s center of gravity (Haury et al. 1980, Yen et al. 1992, Kiørboe et al. 1999). The concept usually applied in this situation is that of water deformation, which can be thought of as the spatiotemporal changes undergone by an initially spherical parcel of water that is distorted into an ellipsoid by the disturbance. Described by the mathematical construct of a deformation tensor (Batchelor 1967), this results in the sphere elongating and narrowing, so for a parcel centered on an organism the relative water flow is outward away from the organism’s center along certain axes and inward along others (see Kiørboe and Visser 1999, for a more detailed treatment). A simplified case that has been used to model an approaching object computes the disturbance produced by a sphere displaced along an axis defining θ = 0 (in spherical coordinates) in water treated as an incompressible inviscid fluid. The maximum water velocity, u, at a distance, r, directly in front of such an object approaching at a speed, U, is given by:

3 u = U ( a / r ) (1)

where a is the radius of the sphere (Kalmijn 1988). This disturbance falls off as the cube of the distance from the center of the source. However, the on-axis deformation rate of the water directly in front of the source is given by:

∆ = −3Ua 3 / r 4 (2)

where Δ, the deformation rate (in inverse time units, i.e., the number of mm/sec of water movement developed per mm of sensor separation from the body) is the spatial derivative of the flow (Kiørboe and Visser 1999). Thus, the deformation rate, which determines the magnitude of the near-field hydrodynamic flow that a pelagic organism can sense, falls off with a steeper, fourthpower dependence on the distance from the source. Further, the ability to sense the deformation depends on the orientation of the sensor mechanism with respect to the deforming water parcel, since between the axes of elongation and shortening, there are axes of no change, along which the disturbance is undetectable. In the case of an approaching sphere, the axis of shortening coincides with that of the approach, while a smaller elongation occurs along perpendicular axes of the oblate spheroid so formed (in the absence of orientation information, it is usual to report sensitivities in terms of the maximum strain rate at the sensor). Using the above simplified model to highlight the difference in detection ability required for a pelagic organism lacking a fixed frame of reference versus that for a benthic one, consider that the water movement at a sensor deployed at a distance ℓ from the main body along the line of approach is in the proportion 3ℓ/r (ℓ 80 μ m, consistent with observation. Capture of motile prey (flagellates



Mechanoreception in Crustaceans of the Pelagic Realm

and ciliates) extended to smaller sizes, correlating with improved detection due to the generation of larger hydrodynamic cues, as computed from prey size and swimming speeds (Kiørboe et al. 2010). Herbivorous feeding by copepods on particles entrained in a feeding current has been typically ascribed to chemosensory input from odor plumes arriving in advance of the algal cell (Poulet and Marsot 1978, Strickler 1982); however, mechanosensory information may be important as well (DeMott and Watson 1991, Bundy et al. 1998, Bundy and Vanderploeg 2002). A detailed hydrodynamic model of the fluid deformations surrounding the herbivore and the algal cell supports the conclusion that for larger phytoplankton cells, mechanoreception alone could explain experimental observations (Bundy and Vanderploeg 2002). Tactile Detection Tactile detection of a potential food (particularly prey) has been studied in the cladoceran Leptodora. In this crustacean, predatory attacks were only triggered after direct contact with a potential prey (Browman et al. 1989). The related fairy shrimp (Branchinecta gigas) inhabits vernal pools of high turbidity. These raptorial predators have unusual, putatively mechanoreceptive and/or chemoreceptive setae to detect their prey in these low-visibility environments (Boudrias and Pires 2002). Food Capture and Processing Once food is captured there is additional sensory input that is critical for the ingestion of the food particle. At this stage, information is gathered to determine the quality of the food, and to process it for ingestion. A combination of mechanosensory and chemosensory input is used to decide whether to ingest or reject the prey item based on (apparently) gustatory information (Paffenhöfer et al. 1982). Communication Mating Behaviors Communication between members of the opposite sex is critical in mate recognition (Buskey 1998). Chemosensory tracking by males has been described for several copepod species (Yen and Lasley 2011). However, mate tracking can also be mediated by mechanosensory information in some copepod species, in particular those with a reduced number of chemosensory setae, such as Acartia spp. (Fig. 11.1C) (Strickler 1998, Bagøien and Kiørboe 2005). Once contact has been made, male-female pairs engage in a “tandem dance,” which appears to be mediated by mechanosensory information, as the male closely follows the females producing tandem jumps prior to mating (Doall et  al. 1998). In brine shrimp (Artemia salina), mechanosensory input enables the synchronized swimming behavior in precopulatory male-female pairs, which reduces energy expended in tandem swimming through phase synchronization (Lent 1977). It has been suggested that this behavior is mediated by putative mechanosensory setae on the antennules (Tyson and Sullivan 1979), but no physiological studies are available. Schooling Behavior Aggregation behaviors, such as swarming and schooling are common in pelagic crustaceans. They are involved in decreasing risk of predation, saving energy, and enhancing reproduction and group foraging (Hamner and Hamner 2000). Mechanosensory information may be critical to maintain interindividual distances in dense aggregations, however

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ORGANIZATION OF CUTICULAR MECHANORECEPTORS Mechanosensilla Anatomical Components Many crustacean cuticular sensilla are located on appendages (Derby 1982), and they are innervated by mechano- and/or chemosensory dendrites (Figs. 11.4 and 11.5). In pelagic crustaceans, the first and second antennae play important roles in distance perception of hydrodynamic and chemical cues (Strickler and Bal 1973, Ball and Cowan 1977, Gill 1985, Lenz et al. 1996), and they display morphologies that are adapted to their pelagic life histories. These appendages extend either anteriorly or laterally from the main axis of the body, presumably to optimize sensory coverage and/or localization of stimuli. The long antennae, which can exceed body length by two- to three-fold, place the setal receptors at a distance from the main body. This increases the space over which f luid deformation can generate a detectable relative motion with respect to the bulk of the organism, while at the same time tending to isolate the sensitive receptors they bear from water movements generated by the swimming appendages (Denton and Gray 1985). These are adaptations well suited to an environment with no fixed point of reference. Setae are typically differentiated along the length of the antennae. In the sergestid shrimp, Acetes sibogae australis, the long second antennae are partitioned by a >90° bend into two regions having different types of setae (Fig. 11.4) (Ball and Cowan 1977). Putatively mechanoreceptive setae in the category designated “type 2” by these authors are found only distal to the bend, while those designated types “3” through “6” are predominantly located proximal to the flexure, and a pair of long “type 7” setae are located at the distal tip. In the calanoid copepods, spiniform setae of the distal tip of the first antennae tend to be especially long and lacking in apical pores, thus appearing adapted for pure mechanoreception. In contrast, those of the antennal shaft possess apical pores (Fig. 11.5), are innervated by two types of dendrites, and thus are likely to be bimodal (Lenz et al. 1996). Setal Form The first filter for information reaching the nervous system is the physical characteristics of the modified cuticular structure: this determines whether any given stimulus will displace the seta (or cuticle) sufficiently to elicit a behavioral response. Crustacean setae vary in length and form from simple conical structures to structures adorned with setules, which in turn can range from scale-like to ribbon-like to plumose. Cuticular sensilla respond to tactile cues, water flow, or bending. As discussed above, the longer hairs may be expected to be sensitive to lower frequencies (more slowly changing water deformations), as will those with more setulation. The geometry of the setal hinge, if present, determines best movement and usually is a good predictor for the plane of highest sensitivity (Douglass and Wilkens 1998), while a stiffer hinge is expected to decrease sensitivity and shift the corner frequency for the filter upward. Bend receptors respond to flexing of the setal shaft (Crouau 1981, Garm et al. 2004).



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Fig. 11.4. Light and scanning electron micrographs of setae on the second antenna of Acetes sibogae australis. (A) Micrograph showing guard (noninnervated) setae (type 1) and two types of mechanosensory setae (type 2A, 2B). (B) SEM image showing the bases of setae types 1 and 2 (numbered labels); und: undetermined setal type. (C) Light micrograph showing how two type 1 setae flank the mechanosensory type 2 seta. From Ball and Cowan 1977, with permission from The Royal Society.

Cellular Organization Setal mechanoreceptors include one or more bipolar sensory cells surrounded by at least three enveloping cells (Fig. 11.6). The dendrites are divided into proximal (= inner) and distal (= outer) dendritic regions, which are separated by the basal body. Both mechano- and chemosensilla have a ciliary origin, and these two types of sensory cells share many features. Five morphological criteria have been identified as establishing mechanosensory function (Altner et al. 1983). The first criterion is presence of a scolopale, an intracellular structure composed of actin filaments (Wolfrum 1990, Weatherby and Lenz 2000) that provides rigid support to the adjacent dendrite(s) (Fig. 11.6C). The presence of a scolopale is the defining feature of a “scolopidial” receptor, which includes the chordotonal organs of insects and crustaceans as well as the majority of crustacean setal receptors, but significantly, not the analogous (if not homologous) insect setal receptors. Characteristic of scolopidial receptors generally are possession of: one to a few sensory neurons, the somata of which are enveloped in a glial cell; a scolopale-containing cell surrounding the sensory dendrite(s); and at least one “attachment cell” at the distal end (Yack 2004). The second criterion is the presence of densely packed microtubules in the distal dendrites (Fig. 11.6B–C). Other criteria include a 9 × 2 + 0 arrangement of microtubules in the ciliary region (Fig. 11.6D) followed proximally by a large ciliary rootlet (Crouau 1982). Connections between the dendritic membrane and the scolopale cell via desmosomes in the vicinity of the

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Fig. 11.5. Scanning electron micrograph of sensory setae on the first antenna of Pleuromamma xiphias adult male (Copepoda). Setae shown include two aesthetascs and two spiniform bimodal setae (arrow heads). Large aesthetascs occur in males only. Scale bar: 43 μm (photo © Tina Weatherby by courtesy and with permission).

basal body constitute the fifth criterion (Altner et al. 1983, Schmidt and Gnatzy 1984). These criteria apply to crustacean mechanoreceptors but not necessarily to those of insects. The relevant stimulus in the insects is thought to be compression of the tubular body (Keil 1997). In the crustaceans, the dendrites are surrounded by an extracellular dendritic sheath distally, and dendrites and sheath become coupled to the cuticle (Ball and Cowan 1977, Schmidt and Gnatzy 1984, Schmidt 1990). Movement of the setae or deformation of the cuticle has been hypothesized to activate the receptor by stretching the dendritic membrane (Crouau 1997). Variations in Design There are no two identical arrangements for the cuticular sensilla, and variations to the basic mechanoreceptor design might provide additional insights into structure-function relationships. Mechanoreceptors can be divided into three functional units: a distal anchor, a proximal anchor, and a stiff intermediate region (Figs. 11.6, 11.7A–C). The nature of the distal and proximal attachments varies, as does the degree of stiffness in the intermediate region. Large rootlets and desmosomes between the membrane of the dendrite and the scolopale cell serve as proximal anchors (Fig. 11.7B–C). In addition to the rootlet, the mysid Antromysis juberthiei has an attachment cell with microtubule bundles that link the proximal dendrites to the cuticle (Fig. 11.7B) (Crouau 1982, 1997). In contrast to this type of anchoring, only small ciliary rootlets have been found in the antennular mechanosensory dendrites of copepods (Weatherby et al. 1994, Weatherby and Lenz 2000) (Figs. 11.6, 11.7A). Instead the scolopale cell is anchored to the cuticle via microtubule bundles located within the first enveloping cell (Figs. 11.6, 11.7A), reminiscent of the ones described for the mysid. However, in the copepod, these attachments occur distal to the basal body region (Fig. 11.7A). The intermediate region, which extends from the basal body to the seta, is characterized by a proliferation of microtubules in the distal dendrites, and the presence of the scolopale

Fig. 11.6. Drawing and transmission electron (TEM) micrographs through a mechanoreceptor on the first antenna of Pleuromamma xiphias (Copepoda). Drawing is based on a longitudinal cross-section through the dendrites and supporting cells. In addition to the scolopale cell (containing the scolopale), there are three enveloping cells. One enveloping cell is located within the first antenna only and it has microtubules that transverse the cell and attach to the cuticle along the anterior antennular edge. The other enveloping cells are more distal and they accompany the distal dendrites into the lumen of the seta. Lines through the drawing indicate approximate locations of the TEM cross sections. (a) attachment of dendrites to setal cuticle; (b) distal dendrites near the base of the seta beyond the scolopale; (c) distal dendrites showing tightly packed microtubules and surrounding scolopale; (d)  basal body region showing dendrites surrounded by liquor cavity and a narrow scolopale. Scale bars: a, 0.5 μm; b, c, and d, 1 μm; insets: b, c and d, 0.1 μm. Abbreviations: bb, basal body; bl, basal lamina; Cu, cuticle; dd, distal dendrites; lc, liquor cavity; mt, microtubules; nu, nucleus; pd, proximal dendrite; sc, scolopale. Figure redrawn from Weatherby et al. 1994 and Weatherby and Lenz 2000.

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Petra H. Lenz and Daniel K. Hartline structure inside of the first enveloping cell. Both the scolopale and the microtubules have a stiffening function. Distal dendrites are densely packed with microtubules, however, the numbers range over two orders of magnitude from 30 to 3,000 among crusraceans (Ball and Cowan 1977, Crouau 1982, Altner et al. 1983, Weatherby et al. 1994). Scolopales vary from poorly developed (first antenna: Balanus amphitrite cyprid; Lagersson et al. 2003) to either a broad crescent (sergestid second antenna: A. sibogae australis; Fig. 11.8C–D) (Ball and Cowan 1977) or even an enclosed tube (Fig. 11.6C; copepod first antenna: Cyclops scutifer, P. xiphias) (Strickler and Bal 1973, Weatherby and Lenz 2000). In addition to the scolopale, tubular sheath cells with an abundance of microtubules add stiffness in A. sibogae australis (Fig. 11.7C). At the proximal end, the scolopale thins (Fig. 11.6D) and terminates at the level of the rootlet (Fig. 11.7A–C). The length of the scolopale differs among setal receptors. In the copepod, it extends to the setal hinge (Fig. 11.7A), while in the sergestid and in the mysid it terminates more proximally (Fig.  11.7B–C). The copepods are characterized by having the best-developed scolopale and the highest densities of microtubules with either one or two mechanosensory dendrites tightly packed within a scolopale tube (Strickler and Bal 1973, Weatherby et al. 1994, Weatherby and Lenz 2000). These dendrites contain microtubules numbering in the thousands packed in a quasi-crystalline array (Fig. 11.6B) (Weatherby et al. 1994). The copepod mechanoreceptors appear to be exceptionally rigid, which undoubtedly contributes to their high mechanical sensitivity and possibly frequency response (see below). In some cases, the scolopale may be lacking entirely, yet other characteristics of the dendrite suggest mechanosensory function. This type of putative mechanoreceptor has been found on the mouthparts of calanoid copepods (Paffenhöfer and Loyd 1999, 2000). Other examples in benthic crustaceans include putative mechanosensory sensilla in isopods (Alexander 1977, Brandt 1988). Sensilla on the first and second antennae of Hutchinsoniella macracantha also lack scolopales, leading Elofsson and Hessler (1991) to conclude an absence of mechanosensors, which seems unlikely. The setae are innervated by multiple dendrites, one with a high density of microtubules, which is often indicative of mechanosensory function. Distal anchoring of individual dendrites occurs either at the base or within the seta along the distal wall in alignment with the plane of movement (Fig. 11.7A–C). At the distal end, dendrites usually terminate within an extracellular dendritic sheath (typically electron-dense), which appears to be firmly attached to the cuticle (Fig. 11.7B–C, 11.8B) (Ball and Cowan 1977, Crouau 1982, Brandt 1988, Schmidt 1990), although in some cases there is no clear evidence of anchoring to the cuticle (Cash-Clark and Martin 1994, Lagersson et al. 2003). In the copepod first antenna, there is a close apposition between dendrites and sheath cells with the cuticle, giving the appearance of a tight connection (Fig. 11.6A, 11.7A) (Weatherby and Lenz 2000). In the antennular palisade seta organ in crayfish, the distal dendrites terminate in a chorda, which is reminiscent of the organization of chordotonal organs (Kouyama and Shimozawa 1982). Bimodal Receptors Many cuticular setae in the Crustacea are bimodal, responding to both chemical and mechanical stimuli (Hallberg and Skog 2011). For example, of the 57 spiniform setae on the first antenna of the calanoid copepod P. xiphias, 48 are mixed modality and nine are mechanosensory only (Lenz et al. 1996). In most cases, sensilla are innervated by unimodal sensory cells, which can be identified as such morphologically (Ball and Cowan 1977, Altner et al. 1983, Schmidt and Gnatzy 1984). Chemosensory dendrites differ from the mechanosensory ones by the presence of fewer microtubules, small ciliary rootlets, and their location within the dendritic sheath (Altner et al. 1983). The dynein arms of the ciliary (9 + 0) microtubule arrangement are not clearly visible in the chemosensory dendrites. Typically mechano- and chemosensory dendrites are bundled



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Fig. 11.7. Schematic drawings of mechanoreceptors from three pelagic crustaceans (A) Pleuromamma xiphias. (B) Antromysis juberthiei. (C) Acetes sibogae australis. Mechanoreceptors in all three species are designed as second order levers. Diagrams based on drawings, electron micrographs and descriptions in Ball and Cowan (1977), Crouau (1982), and Weatherby and Lenz (2000). Drawing not to scale. From www.pbrc. hawaii.edu with permission.

together within the liquor cavity and enclosed by the dendritic sheath distally or the scolopale cell in the intermediate region (Altner et al. 1983, Cate and Derby 2002). In calanoid copepods, the chemosensory dendrites are much shorter than the mechanosensory ones, and their cell bodies are located at the base of the seta (Lenz et al. 1996). Bimodal sensory cells, that is sensory cells that respond to both chemical and mechanical stimuli, have been reported for the walking legs of a crayfish (Austropotamobius torrentium) (Hatt 1986). Bend Receptors Bend receptors have been reported for sensory setae on the mouthparts. A morphological study focused on the feeding appendages of calanoid copepods (Temora stylifera and Centropages velificatus) discovered an unusual arrangement of the distal dendrites within the setae, which led the authors to propose that these mechanosensors may be sensitive to bending (Paffenhöfer and Loyd 1999, 2000). In the calanoids, the putative bend receptors are innervated by both mechanosensory and chemosensory dendrites and they occur on feeding appendages (Paffenhöfer and Loyd 1999, 2000), which is consistent with a role in feeding behavior. In Panulirus argus, bend receptors were identified physiologically on mouthparts (mandibular palp, maxillipeds) (Garm et al. 2004). Simple and cuspidate setae occur on these appendages, and they are innervated by three types of neurons: one chemosensory, one clearly mechanosensory, and the third most

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Fig. 11.8. Micrographs of mechanosensory sensilla of Acetes sibogae australis (A–C) and Neomysis integer (D). (A) Light micrograph of a type 4 seta from the second antenna. Scale: 2µm (B) TEM micrograph of a cross section through a seta type 4 showing the distal end of a sensory dendrite surrounded by the dendritic sheath, which appears to be attached to the cuticle (arrow, dd). (C) TEM micrograph of a cross section through the distal dendrite region showing the scolopale (Sc) partially surrounding a liquor cavity with three sensory dendrites (arrows, dd). (D) TEM micrograph of a cross section through three mechanosensors from three setae on the first antenna. Each seta is innervated with three dendrites located within a liquor cavity and partially surrounded by a scolopale. Two sets of dendrites are filled with microtubules large arrowheads) and third one shows ciliary necklaces (9 + 0) (thin arrowheads). A–C from Ball and Cowan 1977, with permission from The Royal Society, and D from Hallberg and Hansson 1997, with permission from Wiley.

likely mechanosensory (Garm and Høeg 2006). Interestingly, the distal dendrites are characterized by a long dendritic sheath that extends for most of the setal length. The scolopale is located in the appendage itself and limited to the ciliary region.

MECHANOSENSORY MECHANISMS Physiologically, antennular (first antenna) mechanoreceptors of pelagic crustaceans are capable of exceptional performances, sensitive both to very small water movements and high frequencies (Wiese and Marschall 1990, Yen et al. 1992). Basic mechanisms appear to be similar to those in benthic crustaceans and insects, but some pelagic crustaceans have evolved variations that enhance their mechanoreceptive capabilities. The need for enhanced sensitivity in a pelagic environment has been mentioned above, and indeed the literature on mechanoreception stresses the requirement as if it were a significant problem. However, mechanotransduction is intrinsically highly sensitive. Auditory hair cell transduction channels, representative of other stretch-sensitive channels, respond in the nanometer range (Hudspeth and Corey 1977). Movements of this magnitude are not difficult to generate in an aquatic environment, and often the problem may be the reverse—too much sensitivity (Barth 2004). This might be particularly appreciated in insect mechanoreceptors in which the relatively large motion of the tip of a seta is reduced by lever principles to very small movements of dendritic membrane at the tubular body where transduction is presumed to occur. As mentioned above, the insertion of the dendrites in



Mechanoreception in Crustaceans of the Pelagic Realm

copepod antennular setae occurs farther out in a seta, where the motion of the seta is not diminished as greatly as at its base. Extracellular Recordings In copepods, nerve impulses can be recorded extracellularly from the primary mechanoreceptors (Lenz et al. 1989, Gassie et al. 1993, Fields et al. 2002). These take the form of “giant” spikes (mV) with unusual characteristics in amyelinate augaptiloid and centropagoid copepods (Hartline et al. 1996). In these copepods, two reidentifiable mechanoreceptor units can be detected in each antennule (Fig. 11.9). These originate in neuronal somata innervating sensory setae of each distal tip (Hartline et al. 1996). The receptor neurons are capable of firing spikes at rates up to several kHz (Hartline et al. 1996, Fields and Weissburg 2004), an exceptionally high rate for a neuron, and their spike trains can phase-lock to stimuli to about 2 kHz (Lenz 1993, Hartline et al. 1996). In myelinate calanoid groups, extracellularly recorded impulses from antennular mechanoreceptors are much smaller than in amyelinates (Lenz et al. 2000, Funk 2005). Sensitivity Copepod antennular mechanoreceptors are sensitive to very small hydrodynamic disturbances, including abrupt displacements and oscillatory water movements (Fig. 11.9A). The frequency range for oscillatory stimuli extends up to and above 2 kHz, unusually high for aquatic arthropods (Hartline et al. 1996), yet consistent with the prediction that these setae are high-pass filters (Fig. 11.2B). At their best frequency of about 700 Hz, spike thresholds for the giant mechanoreceptor axons of the calanoids P. xiphias and L. madurae are as low as 10 nm of water movement, and responsiveness to oscillatory water velocities is relatively f lat from 100 to in excess of 1,000 Hz (Fig. 11.9A) (Yen et al. 1992, Lenz and Yen 1993, Hartline et al. 1996). Funk’s studies suggest that the small-spike units, too, have high sensitivity to high-frequency water movement, even in early (copepodid) developmental stages (Funk 2005). Threshold sensitivity curves reported for euphausids suggest that these pelagic organisms are sensitive to high frequencies as well (maximum frequency tested:  400 Hz) (Wiese and Marschall 1990, Patria and Wiese 2004). Using a different stimulus, Fields et al. (2002) found that the long stiff seta on the distal tip of Gaussia princeps is highly sensitive and responds at 50% maximum firing rates to angular displacements of 0.9° in the distal direction, while the more f lexible seta was characterized by lower sensitivity (> 5°) (Fields et  al. 2002). Even though these setae are dually innervated (Weatherby and Lenz, unpublished data), the neural discharges are characterized by unitary spikes, suggesting that the two neurons are firing synchronously. Behavioral studies in the mesopelagic P. xiphias and the shallow-water L. madurae show that the same types of disturbances that elicit firing in the mechanosensory neurons can trigger rapid escape reactions (Hartline et al. 1996). The physiological sensitivity measured in the pontellid L. madurae is equal to that for the behavioral escape reaction in the most sensitive range (Hartline et al. 1996). Thus, it may be that even a single nerve impulse can elicit a behavioral reaction in these animals. Myelin and Mechanosensory Triggering of Escape Behavior Morphological studies of the sensory neurons, interneurons, and motor neurons involved in the mechanosensory triggering of the escape reaction have revealed that some, but not all, copepods

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Fig. 11.9. Neural responses to water movement in copepod setal receptors located on the distal tip of the antennules of the calanoid copepod Pleuromamma xiphias. (A) Threshold water velocities as a function of stimulus frequency. Each symbol represents threshold data obtained for different preparations. (B) Extracellular recordings of nerve impulses in response to a hydrodynamic stimulus produced by the displacement of a 3-mm sphere moving parallel to the antennular axis at 200 Hz as shown by the bottom trace (“Displacement”). Water displacement at the tip of the antennule was calculated at 25 nm using dipole equations (Kalmijn 1988). Two large units, “A” and “B” could be identified reliably in this species. Reprinted with permission from www.pbrc.hawaii.edu/lucifer.

possess nerve fibers ensheathed in myelin (Davis et al. 1999, Lenz et al. 2000, Weatherby et al. 2000). The myelinate taxa correspond to those with the very small impulses recorded extracellularly from primary sensory neurons (Lenz et  al. 2000, Funk 2005). Much of the reason for the small amplitude appears to be the restriction of active membrane to small patches of nodal membrane, as in vertebrate myelinated axons. Myelin in vertebrates speeds nerve impulse propagation by an order of magnitude over that of an unmyelinated fiber of the same diameter (Hartline and Colman 2007), allowing a large animal to react rapidly enough to survive. The occurrence of giant fibers in copepods, as well as myelin (and giant nerve fibers of Drosophila for that matter) suggests that every millisecond counts. Myelin potentially shortens the mechanosensory reaction time for a 2- to 3-mm-long copepod by about 2 msec just in reduced conduction time of nerve impulses (Lenz et al. 2000). Compared to about 6 msec for an amyelinate species such as Pleuromamma, this is a significant fraction of an already very fast reaction. Myelinated axons are also present in other pelagic crustaceans including the Dendrobranchiata and the Caridea (Heuser and Doggenweiler 1966, Xu and Terakawa 1999) and some of the fastest nerve



Mechanoreception in Crustaceans of the Pelagic Realm

impulse conduction velocities ever reported are in penaeid shrimp (Xu and Terakawa 1999). Myelination of giant axons has been documented for two other important pelagic crustacean groups:  euphausids and mysids (Hartline and Kong 2008). Among copepods, the presence/ absence of myelin has been correlated with large-scale distribution patterns, which suggest that myelinate taxa may be better at withstanding high predation risk from visual predators (Lenz 2012). Myelination also has the potential for significantly improving the accuracy of localization of the direction of a predatory attack. An important component of the ability of a terrestrial organism to localize a sound source is the difference in arrival time of the sound at two spatially separated auditory receptors (“ears”). This arrival-time difference is much less in aquatic organisms and for near-field disturbances is probably inconsequential for the sensory systems described so far. However, the difference in cue amplitude between antennae, owing to the steep spatial attenuation of near-field water deformation, can be very large, as demonstrated for the sergestid shrimp A. sibogae australis (Denton and Gray 1985). A difference in time-to-threshold of nerve impulses from the two antennae could impart directional information. Rapid conduction could improve the precision of such timing. Such speculation remains to be tested experimentally.

FUTURE DIRECTIONS This review underscores the importance of hydromechanical information to the success of pelagic crustaceans. The lessons learned from such studies are of value in interpreting and understanding other pelagic groups as well, as they are exposed to similar constraints. However, it is clear that much is still to be learned about the sensory environment of pelagic organisms. Perhaps as unfortunate is the spottiness of our information. Much of what we have reviewed here has focused on copepods, a taxon with which the authors have the most experience, but this is not entirely from personal bias. Few recent studies have appeared on the mechanosensory systems of other pelagic crustacean groups, including the ecologically important decapod shrimp, euphausiids, and mysids, let alone larval reptantians. Promising future research directions include much-needed information on the neuroecology of mechanoreception in these groups and more thorough understanding of the physiology of the diverse mechanoreceptors found in pelagic crustaceans. Much of the lack of understanding of mechanoreception in pelagic crustaceans is shared by the better-studied benthic forms. A better understanding of form-function relationships in the design of crustacean mechanoreceptors generally would provide a basis for assessing the type of information that is perceived by both forms. As reviewed, the physics that describes the relative movements of the fluid and a simple conical seta has been characterized fairly well. Less is known about how setules, intersetal spacing and variations in the cuticle as well as the hinge design affect the linkage between fluid movement and displacement of the seta at the transduction site. Transduction is still poorly understood in all crustacean exteroceptors, including localization of the site of transduction, which in turn is dependent on displacement of the seta in response to the hydromechanical stimulus. Given that the crustacean receptors are scolopidial or of scolopidial origin, they may be quite different from the better studied nonscolopidial insect and arachnid setal mechanoreceptors. The best-studied crustacean transduction mechanism is the muscle receptor organ, a receptor with an entirely different (nonciliary) origin from the ciliary exteroceptors of crustaceans and insects. However a wealth of recent information from genetic and molecular approaches has become available as a potential guide. Thus, a comparison of transduction mechanisms between the insect and the crustacean mechanoreceptors might further elucidate function at the molecular level for both.

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SUMMARY AND CONCLUSIONS Pelagic crustaceans lack the substrata-reference of their benthic cousins, hence their setal mechanoreceptors must be sensitive to water deformation rather than to bulk flow. Deformation attenuates more rapidly with distance from a source than does bulk flow, placing greater demands for high-sensitivity sensors. In terms of their physics, setal mechanoreceptors in water act as high-pass filters. This filtering characteristic interacts with physiological processes to determine sensitivity to hydrodynamic cues. As with benthic crustaceans, mechanosensory input is important in predator-prey interactions, feeding behavior, and social behavior. In predator-prey interactions, basic neural circuitry appears similar to that studied in greater depth in benthic Crustacea. Sensitivity to hydrodynamic cues indeed appears to be greater, at least in pelagic calanoid copepods, than in most benthic crustaceans. In the absence of turbulence or habituation, escape responses in calanoid copepods can be triggered with water deformation rates of approximately 1/sec while mechanosensory triggering of bioluminescent discharge on one tested luminescent species was 100 times less sensitive. Predatory pelagic crustaceans use mechanosensory cues to detect potential prey, especially in species lacking well-developed eyes, or under conditions of poor visibility. Mechanoreceptors of pelagic crustaceans have the same basic organization as do those of benthic forms, but may be deployed at substantial distances from the body to enhance sensitivity to water deformation, and may involve internal structures that increase sensitivity to small setal deflections, in particular a well-developed scolopale and large bundles of stiffening microtubules. Physiological recordings, confirmed by some behavioral studies, show evidence of receptor sensitivity to significantly higher frequencies than for benthic forms, presumed to relate to greater sensitivity to rapidly rising water deformations, and consistent with a high-pass filtering characteristic. Several pelagic crustacean taxa possess myelinated axons in their nervous systems, especially in axons thought to participate in rapid escape responses. This appears to have been lost in benthic forms, which in combination with ecological correlates, enhances the impression of high predation risk for pelagic taxa, which lack the hiding places of their benthic relatives. Thus, mechanoreception in pelagic crustaceans employs the same basic hardware used by benthic forms, but evolution has refined it to facilitate adaptation to the particular ecological conditions inherent in a pelagic life. However, many of the details of this refinement have yet to be worked out, from the hydrodynamics of receptor coupling and sensory transduction to the central processing of time-varying 3D sensory landscapes.

ACKNOWLEDGMENTS We thank Cheryl Clarke, Steve Haddock, Tina Weatherby, Ben Clauberg, and Jenn Kong for providing images for this review. We thank Dr. Friedrich Barth for his constructive suggestions to improve an earlier version of the discussion on setal hydrodynamics. Partially supported by funding from NSF 09-23692 (PIs: DKH, PHL and A. Castelfranco).

REFERENCES Alcaraz, M., and J.R. Strickler. 1988. Locomotion in copepods—pattern of movements and energetics of Cyclops. Hydrobiologia 167:409–414. Alcaraz, M., G.A. Paffenhöfer, and J.R. Strickler. 1980. Catching the algae: a first account of visual observations on filter-feeding calanoids. Pages 241–248 in W.C. Kerfoot, editor. Evolution and ecology of zooplankton communities. University Press of New England, Hanover, NH.



Mechanoreception in Crustaceans of the Pelagic Realm

Alexander, C.G. 1977. Antennal sense organs in the isopod Ligia oceanica (Linn). Marine Behaviour and Physiology 5:61–77. Altner, I., H. Hatt, and H. Altner. 1983. Structural properties of bimodal chemosensitive and mechanosensitive setae on the pereiopod chelae of the crayfish, Austropotamobius torrentium. Cell and Tissue Research 228:357–374. Árnadóttir, J., and M. Chalfie. 2010. Eukaryotic mechanosensitive channels. Annual Review of Biophysics 39:111–137. Arnott, S.A., D.M. Neil, and A.D. Ansell. 1999. Escape trajectories of the brown shrimp Crangon crangon, and a theoretical consideration of initial escape angles from predators. Journal of Experimental Biology 202:193–209. Bagøien, E., and T. Kiørboe. 2005. Blind dating—mate finding in planktonic copepods. I. Tracking the pheromone trail of Centropages typicus. Marine Ecology Progress Series 300:105–115. Ball, E.E., and A.N. Cowan. 1977. Ultrastructure of the antennal sensilla of Acetes (Crustacea, Decapoda, Natantia, Sergestidae). Philosophical Transactions of the Royal Society of London B: Biological Sciences 277:429–457. Barth, F.G. 2004. Spider mechanoreceptors. Current Opinion in Neurobiology 14:415–422. Batchelor, G.K. 1967. An introduction to fluid dynamics. Cambridge University Press, Cambridge. Bauer, R.T. 2004. Remarkable shrimps: adaptations and natural history of the carideans. University of Oklahoma Press, Norman. Boudrias, M.A., and J. Pires. 2002. Unusual sensory setae of the raptorial Branchinecta gigas (Branchiopoda: Anostraca). Hydrobiologia 486:19–27. Bradley, C.J. 2009. Development of copepod escape behaviors in response to a hydrodynamic stimulus. MS thesis, University of Hawaii at Manoa. Brandt, A. 1988. Morphology and ultrastructure of the sensory spine, a presumed mechanoreceptor of Sphaeroma hookeri (Crustacea, Isopoda), and remarks on similar spines in other peracarids. Journal of Morphology 198:219–229. Browman, H.I., S. Kruse, and W.J. O’Brien. 1989. Foraging behavior of the predaceous cladoceran, Leptodora kindti, and escape responses of their prey. Journal of Plankton Research 11:1075–1088. Browman, H.I., J. Yen, D.M. Fields, J.-F. St-Pierre, and A.B. Skiftesvik. 2011. Fine-scale observations of the predatory behaviour of the carnivorous copepod Paraeuchaeta norvegica and the escape responses of their ichthyoplankton prey, Atlantic cod (Gadus morhua). Marine Biology 158:2653–2660. Bundy, M.H., and H.A. Vanderploeg. 2002. Detection and capture of inert particles by calanoid copepods: the role of the feeding current. Journal of Plankton Research 24:215–223. Bundy, M.H., T.F. Gross, H.A. Vanderploeg, and J.R. Strickler. 1998. Perception of inert particles by calanoid copepods: behavioral observations and a numerical model. Journal of Plankton Research 20:2129–2152. Burdick, D.S., D.K. Hartline, and P.H. Lenz. 2007. Escape strategies in co-occurring calanoid copepods. Limnology and Oceanography 52:2373–2385. Bush, B.M.H., and M.S. Laverack. 1982. Mechanoreception. Pages 399–468 in H. L. Atwood and D.C. Sandeman, editors. The Biology of Crustacea. Vol. 3. Neurobiology: structure and function. Academic Press, New York. Buskey, E.J. 1998. Components of mating behavior in planktonic copepods. Journal of Marine Systems 15:13–21. Buskey, E.J., P.H. Lenz, and D.K. Hartline. 2012. Sensory perception, neurobiology and behavioral adaptations for predator avoidance in planktonic copepods. Adaptive Behavior 20:57–66. Cash-Clark, C.E., and J.W. Martin. 1994. Ultrastructure of the frontal sensory fields in the Lynceidae (Crustacea, Branchiopoda, Laevicaudata). Journal of Morphology 221:153–160. Cate, H.S., and C.D. Derby. 2002. Ultrastructure and physiology of the hooded sensillum, a bimodal chemo-mechanosensillum of lobsters. Journal of Comparative Neurology 442:293–307. Chen, D.S., G. Van Dykhuizen, J. Hodge, and W.F. Gilly. 1996. Ontogeny of copepod predation in juvenile squid (Loligo opalescens). Biological Bulletin 190:69–81. Clarke, R.D., E.J. Buskey, and K.C. Marsden. 2005. Effects of water motion and prey behavior on zooplankton capture by two coral reef fishes. Marine Biology 146:1145–1155.

315

316

Petra H. Lenz and Daniel K. Hartline Coombs, S. 1994. Nearfield detection of dipole sources by the goldfish (Carassius auratus) and the mottled sculpin (Cottus bairdi). Journal of Experimental Biology 190:109–129. Crouau, Y. 1981. Cytology of various antennal setae in a troglobitic Mysidacea (Crustacea). Zoomorphology 98:121–134. Crouau, Y. 1982. Primary stages in the sensory mechanism of the setulate sensilla, external mechanoreceptors of a cavernicolous Mysidacea. Biology of the Cell 44:45–56. Crouau, Y. 1997. Comparison of crustacean and insect mechanoreceptive setae. International Journal of Insect Morphology and Embryology 26:181–190. Davis, A.D., T.M. Weatherby, D.K. Hartline, and P.H. Lenz. 1999. Myelin-like sheaths in copepod axons. Nature 398:571–571. Dechant, H.E., F.G. Rammerstorfer, and F.G. Barth. 2001. Arthropod touch reception: stimulus transformation and finite element model of spider tactile hairs Journal of Comparative Physiology A 187:313–322, erratum in 851. DeMott, W.R., and M.D. Watson. 1991. Remote detection of algae by copepods—responses to algal size, odors and motility. Journal of Plankton Research 13:1203–1222. Denton, E.J., and J. Gray. 1985. Lateral-line-like antennae of certain of the Penaeidea (Crustacea, Decapoda, Natantia). Proceedings of the Royal Society of London B: Biological Sciences 226:249–261. Derby, C.D. 1982. Structure and function of cuticular sensilla of the lobster Homarus americanus. Journal of Crustacean Biology 2:1–21. Devarakonda, R., F.G. Barth, and J.A.C. Humphrey. 1996. Dynamics of arthropod filiform hairs. IV. Hair motion in air and water. Philosophical Transactions of the Royal Society of London B: Biological Sciences 351:933–946. Doall, M.H., S.P. Colin, J.R. Strickler, and J. Yen. 1998. Locating a mate in 3D: the case of Temora longicornis. Philosophical Transactions of the Royal Society of London B: Biological Sciences 353:681–689. Doall, M.H., J.R. Strickler, D.M. Fields, and J. Yen. 2002. Mapping the free-swimming attack volume of a planktonic copepod, Euchaeta rimana. Marine Biology 140:871–879. Douglass, J.K., and L.A. Wilkens. 1998. Directional selectivities of near-field filiform hair mechanoreceptors on the crayfish tailfan (Crustacea: Decapoda). Journal of Comparative Physiology A 183:23–34. Drenner, R.W., J.R. Strickler, and W.J. O’Brien. 1978. Capture probability: the role of zooplankter escape in the selective feeding of planktivorous fish. Journal of the Fisheries Research Board of Canada 35:1370–1373. Elofsson, R., and R.R. Hessler. 1991. Sensory morphology in the antennae of the cephalocarid Hutchinsoniella macracantha. Journal of Crustacean Biology 11:345–355. Fields, D.M., and M.J. Weissburg. 2004. Rapid firing rates from mechanosensory neurons in copepod antennules. Journal of Comparative Physiology A 190:877–882. Fields, D.M., and J. Yen. 1996. The escape behavior of Pleuromamma xiphias in response to a quantifiable fluid mechanical disturbance. Pages 323–339 in P.H. Lenz, D.K. Hartline, J.E. Purcell, and D.L. Macmillan, editors. Zooplankton: sensory ecology and physiology. Gordon and Breach Publishers, Amsterdam. Fields, D.M., and J. Yen. 2002. Fluid mechanosensory stimulation of behavior from a planktonic marine copepod Euchaeta rimana Bradford. Journal of Plankton Research 24:747–755. Fields, D.M., D.S. Shaeffer, and M.J. Weissburg. 2002. Mechanical and neural responses from the mechanosensory hairs on the antennule of Gaussia princeps. Marine Ecology Progress Series 227:173–186. Funk, K.R. 2005. Development of mechanosensory responses in myelinated copepods. MS thesis, University of Hawaii at Manoa, Honolulu. Garm, A. 2004. Mechanical functions of setae from the mouth apparatus of seven species of decapod crustaceans. Journal of Morphology 260:85–100. Garm, A., and J.T. Høeg. 2006. Ultrastructure and functional organization of mouthpart sensory setae of the spiny lobster Panulirus argus: new features of putative mechanoreceptors. Journal of Morphology 267:464–476.



Mechanoreception in Crustaceans of the Pelagic Realm

Garm, A., C.D. Derby, and J.T. Høeg. 2004. Mechanosensory neurons with bend- and osmo-sensitivity in mouthpart setae from the spiny lobster Panulirus argus. Biological Bulletin 207:195–208. Gassie, D.V., P.H. Lenz, J. Yen, and D.K. Hartline. 1993. Mechanoreception in zooplankton 1st antennae—electrophysiological techniques. Bulletin of Marine Science 53:96–105. Gemmell, B.J. 2011. Evasion from predation: the perilous life of planktonic copepods throughout development. PhD dissertation, University of Texas, Austin. Gill, C.W. 1985. The response of a restrained copepod to tactile stimulation. Marine Ecology Progress Series 21:121–125. Goodman, M.B., E.A. Lumpkin, A. Ricci, W.D. Tracey, M. Kernan, and T. Nicolson. 2004. Molecules and mechanisms of mechanotransduction. Journal of Neuroscience 24:9220–9222. Haddock, S.H.D., M.A. Moline, and J.F. Case. 2010. Bioluminescence in the sea. Annual Review of Marine Science 2:443–493. Hallberg, E., and M. Skog. 2011. Chemosensory sensilla in crustaceans. Pages 103–121 in T. Breithaupt and M. Thiel, editors. Chemical communication in crustaceans. Springer, Heidelberg. Hamner, W.M., and P.P. Hamner. 2000. Behavior of Antarctic krill (Euphausia superba): schooling, foraging, and antipredatory behavior. Canadian Journal of Fisheries and Aquatic Sciences 57:192–202. Hartline, D.K., and D.R. Colman. 2007. Rapid conduction and the evolution of giant axons and myelinated fibers. Current Biology 17:R29–R35. Hartline, D.K., and J.H. Kong. 2008. The gain and loss of myelin in malacostracan crustaceans. Program No. 79.22/PP10, Neuroscience, Annual Meeting. Society for Neuroscience, Washington, DC. Hartline, D.K., and P.H. Lenz. 2014. Hydrodynamics of arthropod cuticular setae in water. www.pbrc. hawaii.edu/~danh/seta-hydrodynamics/ accessed February 4, 2014. Hartline, D.K., P.H. Lenz, and C.M. Herren. 1996. Physiological and behavioral studies of escape responses in calanoid copepods. Marine and Freshwater Behaviour and Physiology 27:199–212. Hartline, D.K., E.J. Buskey, and P.H. Lenz. 1999. Rapid jumps and bioluminescence elicited by controlled hydrodynamic stimuli in a mesopelagic copepod, Pleuromamma xiphias. Biological Bulletin 197:132–143. Hatt, H. 1986. Responses of a bimodal neuron (chemo-sensitive and vibration-sensitive) on the walking legs of the crayfish. Journal of Comparative Physiology A 159:611–617. Haury, L.R., D.E. Kenyon, and J.R. Brooks. 1980. Experimental evaluation of the avoidance reaction of Calanus finmarchicus. Journal of Plankton Research 2:187–202. Heuch, P.A., M.H. Doall, and J. Yen. 2007. Water flow around a fish mimic attracts a parasitic and deters a planktonic copepod. Journal of Plankton Research 29:i3–i16. Heuser, J.E., and C.F. Doggenweiler. 1966. The fine structural organization of nerve fibers, sheaths, and glial cells in the prawn, Palaemonetes vulgaris. Journal of Cell Biology 30:381–403. Hudspeth, A.J., and D.P. Corey. 1977. Sensitivity, polarity, and conductance change in the response of vertebrate hair cells to controlled mechanical stimuli. Proceedings of the National Academy of Sciences of the United States of America 74:2407–2411. Humphrey, J.A.C., R. Devarakonda, I. Iglesias, and F.G. Barth. 1993. Dynamics of arthropod filiform hairs. I. Mathematical-modeling of the hair and air motions. Philosophical Transactions of the Royal Society of London B: Biological Sciences 340:423–444. Humphrey, J.A.C., R. Devarakonda, I. Iglesias, and F.G. Barth. 1997. Dynamics of arthropod filiform hairs. I. Mathematical modelling of the hair and air motions. Philosophical Transactions of the Royal Society of London B: Biological Sciences 352:1995 (errata). Humphrey, J.A.C., F.G. Barth, and K. Voss. 2001. The motion-sensing hairs of arthropods: using physics to understand sensory ecology and adaptive evolution. Pages 105–125 in F.G. Barth and A. Schmid, editors. Ecology of sensing. Springer, Berlin. Hwang, J.S., J.H. Costello, and J.R. Strickler. 1994. Copepod grazing in turbulent-flow—elevated foraging behavior and habituation of escape responses. Journal of Plankton Research 16:421–431. Jackson, J.M. 2011. Larval clownfish Amphiprion ocellaris predatory success and selectivity when preying on the calanoid copepod Parvocalanus crassirostris. MS thesis, University of Hawaii at Manoa, Honolulu.

317

318

Petra H. Lenz and Daniel K. Hartline Jiménez, J. 1997. Oceanic turbulence at millimeter scales. Scientia Marina 61:47–56. Johnson, G.E. 1924. Giant nerve fibres in crustaceans, with special reference to Cambarus and Palaemonetes. Journal of Comparative Neurology 36:323–373. Jonsson, A., T.G. Nielsen, I. Hrubenja, M. Maar, and J.K. Petersen. 2009. Eating your competitor: functional triangle between turbulence, copepod escape behavior and predation from mussels. Marine Ecology Progress Series 376:143–151. Jonsson, P.R., and P. Tiselius. 1990. Feeding behavior, prey detection and capture efficiency of the copepod Acartia tonsa feeding on planktonic ciliates. Marine Ecology Progress Series 60:35–44. Kalmijn, A.J. 1988. Hydrodynamic and acoustic field detection. Pages 83–130 in J. Atema, R.R. Fay, A.N. Popper, and W.N. Tavolga, editors. Sensory biology of aquatic animals. Springer, New York. Keil, T.A. 1997. Functional morphology of insect mechanoreceptors. Microscopy Research and Technique 39:506–531. Kils, U. 1992. The ecoSCOPE and dynIMAGE: microscale tools for in situ studies of predator-prey interactions. Archiv für Hydrobiologie, Beihefte 36:83–96. Kiørboe, T., and A.W. Visser. 1999. Predator and prey perception in copepods due to hydromechanical signals. Marine Ecology Progress Series 179:81–95. Kiørboe, T., E. Saiz, and A. Visser. 1999. Hydrodynamic signal perception in the copepod Acartia tonsa. Marine Ecology Progress Series 179:97–111. Kiørboe, T., H.S. Jiang, and S.P. Colin. 2010. Danger of zooplankton feeding: the fluid signal generated by ambush-feeding copepods. Proceedings of the Royal Society of London B: Biological Sciences 277:3229–3237. Kouyama, N., and T. Shimozawa. 1982. The structure of a hair mechanoreceptor in the antennule of crayfish (Crustacea). Cell and Tissue Research 226:565–578. Lagersson, N.C., A. Garm, and J.T. Høeg. 2003. Notes on the ultrastructure of the setae on the fourth antennulary segment of the Balanus amphitrite cyprid (Crustacea: Cirripedia: Thoracica). Journal of the Marine Biological Association of the United Kingdom 83:361–365. Landry, M.D. 1978. Predatory feeding behavior of a marine copepod, Labidocera trispinosa. Limnology and Oceanography 23:1103–1113. Lent, C.M. 1977. The mechanism for coordinating metachronal limb movements between joined male and female Artemia salina during precopulatory behaviour. Journal of Experimental Biology 66:127–140. Lenz, P.H. 1993. Vibration sensitivity in marine copepods. Pages 423–429 in K. Wiese, F.G. Gribakin, A.V. Popov, and G. Renninger, editors. Sensory systems of arthropods. Birkhäuser, Basel. Lenz, P.H. 2012. The biogeography and ecology of myelin in marine copepods. Journal of Plankton Research 34:575–645. Lenz, P.H., and D.K. Hartline. 1999. Reaction times and force production during escape behavior of a calanoid copepod, Undinula vulgaris. Marine Biology 133:249–258. Lenz, P.H., and J. Yen. 1993. Distal setal mechanoreceptors of the first antennae of marine copepods. Bulletin of Marine Science 53:170–179. Lenz, P.H., J. Yen, and D.K. Hartline. 1989. Mechanoreception in the first antenna of marine copepods. Society for Neuroscience Abstracts 15:1288. Lenz, P.H., T.M. Weatherby, W. Weber, and K.K. Wong. 1996. Sensory specialization along the first antenna of a calanoid copepod, Pleuromamma xiphias (Crustacea). Marine and Freshwater Behaviour and Physiology 27:213–221. Lenz, P.H., D.K. Hartline, and A.D. Davis. 2000. The need for speed. I. Fast reactions and myelinated axons in copepods. Journal of Comparative Physiology A 186:337–345. Lowe, E. 1935. On the anatomy of a marine copepod Calanus finmarchicus (Gunnerus). Transactions of the Royal Society Edinburgh 58:561–603. Marrasé, C., E. Saiz, and J.M. Redondo. 1997. Lectures on plankton and turbulence—foreword. Scientia Marina 61:U3. Morris, M.J., G. Gust, and J.J. Torres. 1985. Propulsion efficiency and cost of transport for copepods—a hydromechanical model of crustacean swimming. Marine Biology 86:283–295. O’Brien, W.J. 1987. Planktivory by freshwater fish: thrust and parry in the pelagia. Pages 3–16 in W.C. Kerfoot and A. Sih, editors. Predation direct and indirect impacts on aquatic communities. University Press of New England, Hanover, NH.



Mechanoreception in Crustaceans of the Pelagic Realm

Paffenhöfer, G.A., and P.A. Loyd. 1999. Ultrastructure of setae of the maxilliped of the marine planktonic copepod Temora stylifera. Marine Ecology Progress Series 178:101–107. Paffenhöfer, G.A., and P.A. Loyd. 2000. Ultrastructure of cephalic appendage setae of marine planktonic copepods. Marine Ecology Progress Series 203:171–180. Paffenhöfer, G.A., J.R. Strickler, and M. Alcaraz. 1982. Suspension feeding by herbivorous calanoid copepods—a cinematographic study. Marine Biology 67:193–199. Park, T.S. 1966. The biology of a calanoid copepod Epilabidocera amphitrites McMurrich. Cellule 66:137–244. Patria, M.P., and K. Wiese. 2004. Swimming in formation in krill (Euphausiacea), a hypothesis: dynamics of the flow field, properties of antennular sensor systems and a sensory-motor link. Journal of Plankton Research 26:1315–1325. Poulet, S.A., and P. Marsot. 1978. Chemosensory grazing by marine calanoid copepods (Arthropoda: Crustacea). Science 200:1403–1405. Price, H.J., G.A. Paffenhöfer, and J.R. Strickler. 1983. Modes of cell capture in calanoid copepods. Limnology and Oceanography 28:116–123. Saiz, E., M. Alcaraz, and G.A. Paffenhöfer. 1992. Effects of small-scale turbulence on feeding rate and gross-growth efficiency of three Acartia species (Copepoda, Calanoida). Journal of Plankton Research 14:1085–1097. Schmidt, M. 1990. Ultrastructure of a possible new type of crustacean cuticular strain receptor in Carcinus maenas (Crustacea, Decapoda). Journal of Morphology 204:335–344. Schmidt, M., and W. Gnatzy. 1984. Are the funnel-canal organs the “campaniform sensilla” of the shore crab, Carcinus maenas (Decapoda, Crustacea)? II. Ultrastructure. Cell and Tissue Research 237:81–93. Singarajah, K.V. 1969. Escape reactions of zooplankton: the avoidance of a pursuing siphon tube. Journal of Experimental Marine Biology and Ecology 3:171–178. Strickler, J.R. 1975. Swimming of planktonic Cyclops species (Copepoda: Crustacea): pattern, movements and their control. Pages 599–613 in T. Y.-T. Wu, C.J. Brokaw, and C. Brennan, editors. Swimming and flying in nature. Plenum Press, New York. Strickler, J.R. 1982. Calanoid copepods, feeding currents, and the role of gravity. Science 218:158–160. Strickler, J.R. 1998. Observing free-swimming copepods mating. Philosophical Transactions of the Royal Society B Biological Sciences 353:671–680. Strickler, J.R., and A.K. Bal. 1973. Setae of the first antennae of the copepod Cyclops scutifer (Sars): their structure and importance. Proceedings of the National Academy of Sciences of the United States of America 70:2656–2659. Svetlichnyy, L.S. 1987. Speed, force and energy expenditure in the movement of copepods. Oceanology 27:497–502. Tautz, J. 1979. Reception of particle oscillation in a medium—an unorthodox sensory capacity. Naturwissenschaften 66:452–461. Trager, G., Y. Achituv, and A. Genin. 1994. Effects of prey escape ability, flow speed, and predator feeding mode on aooplankton capture by barnacles. Marine Biology 120:251–259. Tyson, G.E., and M.L. Sullivan. 1979. Antennular sensilla of the brine shrimp, Artemia salina. Biological Bulletin 156:382–392. Viherluoto, M., and M. Viitasalo. 2001. Temporal variability in functional responses and prey selectivity of the pelagic mysid, Mysis mixta, in natural prey assemblages. Marine Biology 138:575–583. Viitasalo, M., T. Kiørboe, J. Flinkman, L.W. Pedersen, and A.W. Visser. 1998. Predation vulnerability of planktonic copepods: consequences of predator foraging strategies and prey sensory abilities. Marine Ecology Progress Series 175:129–142. Viitasalo, M., J. Flinkman, and M. Viherluoto. 2001. Zooplanktivory in the Baltic Sea: a comparison of prey selectivity by Clupea harengus and Mysis mixta, with reference to prey escape reactions. Marine Ecology Progress Series 216:191–200. Visser, A.W. 2001. Hydromechanical signals in the plankton. Marine Ecology Progress Series 222:1–24.

319

320

Petra H. Lenz and Daniel K. Hartline Visser, A.W., H. Saito, E. Saiz, and T. Kiørboe. 2001. Observations of copepod feeding and vertical distribution under natural turbulent conditions in the North Sea. Marine Biology 138:1011–1019. Waggett, R.J., and E.J. Buskey. 2007a. Calanoid copepod escape behavior in response to a visual predator. Marine Biology 150:599–607. Waggett, R.J., and E.J. Buskey. 2007b. Copepod escape behavior in non-turbulent and turbulent hydrodynamic regimes. Marine Ecology Progress Series 334:193–198. Weatherby, T.M., and P.H. Lenz. 2000. Mechanoreceptors in calanoid copepods: designed for high sensitivity. Arthropod Structure and Development 29:275–288. Weatherby, T.M., K.K. Wong, and P.H. Lenz. 1994. Fine-structure of the distal sensory setae on the first antennae of Pleuromamma xiphias Giesbrecht (Copepoda). Journal of Crustacean Biology 14:670–685. Weatherby, T.M., A.D. Davis, D.K. Hartline, and P.H. Lenz. 2000. The need for speed. II. Myelin in calanoid copepods. Journal of Comparative Physiology A 186:347–357. Whitear, M. 1962. The fine structure of crustacean proprioceptors. I. The chordotonal organs in the legs of the shore crab, Carcinus maenas. Philosophical Transactions of the Royal Society London B: Biological Sciences 245:291–324. Widder, E.A. 2010. Bioluminescence in the ocean: origins of biological, chemical, and ecological diversity. Science 328:704–708. Wiese, K., and H.P. Marschall. 1990. Sensitivity to vibration and turbulence of water in context with schooling in Antarctic krill Euphausia superba. Pages 121–130 in K. Wiese, W.D. Krenz, J. Tautz, H. Reichert, and B. Mulloney, editors. Frontiers in crustacean neurobiology. Birkhäuser, Basel. Wilson, C.H., and D.K. Hartline. 2011. The novel organization and development of copepod myelin. I. Ontogeny. Journal of Comparative Neurology 519:3259–3280. Wine, J.J., and F.B. Krasne. 1982. The cellular organization of crayfish escape behavior. Pages 242–292 in D.C. Sandeman and H.L. Atwood, editors. The biology of Crustacea, Vol. 4, Neural integration and behavior. Academic Press, New York. Wolfrum, U. 1990. Actin-filaments—the main components of the scolopale in insect sensilla. Cell and Tissue Research 261:85–96. Wright, D.I., and W.J. O’Brien. 1984. The development and field-test of a tactical model of the planktivorous feeding of white crappie (Pomoxis annularis). Ecological Monographs 54:65–98. Xu, K., and S. Terakawa. 1999. Fenestration nodes and the wide submyelinic space from the basis for the unusually fast impulse conduction of shrimp myelinated axons. Journal of Experimental Biology 202:1979–1989. Yack, J.E. 2004. The structure and function of auditory chordotonal organs in insects. Microscopy Research and Technique 63:315–337. Yen, J., and D.M. Fields. 1992. Escape responses of Acartia hudsonica (Copepoda) nauplii from the flow field of Temora longicornis (Copepoda). Archives of Hydrobiology 36:123–134. Yen, J., and R. Lasley. 2011. Chemical communication between copepods. Pages 177–197 in T. Breithaupt and M. Thiel, editors. Chemical communication in crustaceans. Springer, Heidelberg. Yen, J., P.H. Lenz, D.V. Gassie, and D.K. Hartline. 1992. Mechanoreception in marine copepods— electrophysiological studies on the first antennae. Journal of Plankton Research 14:495–512.

12 THE GEOMAGNETIC SENSE OF CRUSTACEANS AND ITS USE IN ORIENTATION AND NAVIGATION

Kenneth J. Lohmann and David A. Ernst

Abstract Most crustaceans are active, mobile organisms that periodically move through complex environments in search of food, shelter, and mates. In many cases, movements are not random, but instead are directed toward specific targets. On a small spatial scale, destinations can be burrows or other refuges, such as wetter or drier microhabitats; on a larger scale, targets can include locations suitable for larval release or offshore habitats for overwintering. Efficient movement between ecologically important locations has presumably been favored by natural selection, and crustaceans have evolved a suite of guidance mechanisms suitable for directing the movements they make. Among the numerous sensory cues potentially available to terrestrial and aquatic crustaceans, the Earth’s magnetic field is a particularly pervasive cue that is continuously available in nearly all environments that crustaceans inhabit. At least some crustaceans have evolved the ability to perceive the Earth’s field. Various amphipods, isopods, and spiny lobsters possess magnetic compasses, which enable them to maintain headings toward particular directions such as north or south. In principle, the Earth’s field also provides a potential source of positional or “map” information that can help an animal navigate toward a specific target area. The Caribbean spiny lobster, Panulirus argus, has been shown to possess such a magnetic map, which allows lobsters to determine their geographic location relative to a goal. Although behavioral studies have demonstrated that some crustaceans perceive the Earth’s magnetic field, little is known about how they do so. At present, three main mechanisms of magnetoreception have been proposed: electromagnetic induction, magnetite, and chemical magnetoreception. Further studies are needed to determine how widespread magnetoreception is among crustaceans and to investigate the physical basis of the magnetic sense.

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INTRODUCTION The remarkable diversity of crustacean lifestyles is accompanied by extreme variation in movement patterns. A  few species are sedentary, epitomized by barnacles that spend their entire adult lives irreversibly anchored to a hard substratum in a single location. Most crustaceans are mobile to varying degrees, but patterns of movement, as well as the spatial scales traveled, vary greatly among different groups and species. For example, some crustaceans migrate up and down the beach with the tides (Cubit 1969, Forward et  al. 2005, Scapini 2006). Others, such as semiterrestrial crabs, forage over relatively small distances from a burrow but retreat to it when approached by predators (Hughes 1966, Vannini and Cannicci 1995, Zeil 1998). Still others undergo long-distance migrations of considerable duration and complexity (Herrnkind and Kanciruk 1978, Adamczewska and Morris 1998, Tankersley et al. 1998). In all of these cases, survival is enhanced by an ability to orient movements reliably and efficiently toward destinations, regardless of whether the goal is a moist patch of sand, a burrow, or a distant area used for overwintering or larval release. Mobile marine animals exploit numerous types of sensory information while migrating, homing, or moving around their habitats (Lohmann et al. 2008). Among these, the Earth’s magnetic field is unusually reliable and pervasive. In contrast with most other sensory cues, the field is present both night and day, is largely unaffected by weather and season, and exists virtually everywhere in the marine environment, from salt marshes and maritime forests to the deepest ocean trenches. Thus, it is perhaps not surprising that some crustaceans have evolved ways to exploit the geomagnetic field to guide their movements. In this chapter, we review what is known about how crustaceans perceive the Earth’s field and use it in orientation and navigation.

THE EARTH’S MAGNETIC FIELD To a first approximation, the Earth’s magnetic field resembles the dipole field of a giant bar magnet (Fig. 12.1). Field lines leave the southern hemisphere and curve around the globe before reentering the planet in the northern hemisphere. Animals can potentially extract at least two different kinds of information from the Earth’s field. The simplest is directional information, which enables an animal to maintain a consistent heading (e.g., toward the north or south). Animals with this ability are said to have a magnetic compass. Some animals also derive positional information from the Earth’s field; in other words, they can use magnetic cues to assess where they are located relative to a goal, or to determine what direction to travel at a particular location along a complex migratory route (Lohmann et al. 2007, 2012). Animals that derive positional information from the field are said to have a magnetic map. This term is used as convenient shorthand and does not imply that the map is necessarily detailed or organized in the same way as a human map (Lohmann 2010). We will begin by looking at how crustaceans use directional information in the Earth’s magnetic field. Magnetic compasses have been shown to exist in amphipods, isopods, and spiny lobsters.

A MAGNETIC COMPASS IN AMPHIPODS AND ISOPODS Amphipods, also known as sandhoppers or beachhoppers, are common inhabitants of intertidal zones. Many species move with the tides to remain in damp sand, following the receding water to avoid desiccation as the tide falls, but retreating before the rising tide to avoid inundation



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Fig. 12.1. (A) Diagram of the Earth’s magnetic field, illustrating how field lines (represented by arrows) intersect the Earth’s surface, and how inclination angle (the angle formed between the field lines and the Earth) varies with latitude. At the magnetic equator (the curving line across the Earth), field lines are parallel to the Earth’s surface. The field lines become progressively steeper as one travels north toward the magnetic pole, where the field lines are directed straight down into the Earth and the inclination angle is 90 degrees. (B) Diagram illustrating four elements of geomagnetic field vectors that might, in principle, provide animals with positional information. The field present at each location on Earth can be described in terms of a total field intensity and an inclination angle. The total intensity of the field can be resolved into two vector components: the horizontal field intensity and the vertical field intensity. Whether animals are able to resolve the total field into vector components is not known. Diagram is from Lohmann et al. 2007, with permission from The Company of Biologists.

(Scapini 2006). This type of orientation, in which an animal moves mainly along an axis perpendicular to a waterline, is often referred to as Y-axis orientation (Wiltschko and Wiltschko 1995a). The orientation cues that underlie Y-axis orientation in amphipods have been studied extensively and include several different kinds of environmental information, including visual landscape cues, sun compass orientation, and beach slope (e.g., Pardi and Scapini 1983, Pardi and Ercolini 1986, Ugolini et al. 1988). Early attempts to demonstrate magnetic orientation in amphipods produced contradictory results. Experiments carried out in coastal areas of several European countries indicated that the amphipod Talitrus saltator, when tested in complete darkness, was able to orient nonrandomly (Van den Bercken et al. 1967). The Earth’s magnetic field was considered as a possible orientation cue, but an initial attempt to disrupt the orientation by canceling the ambient field was unsuccessful. In a subsequent study conducted in Italy, Ercolini and Scapini (1972) were unable to replicate the finding of nonrandom orientation in darkness. Working in The Netherlands, however, Arendse (1978, 1980, Arendse and Kruyswijk 1981) reported orientation of Talitrus in directions coinciding with the land-sea axis. When the ambient field was rotated to a new position, the orientation shifted accordingly; when the ambient field was eliminated, the orientation vanished. These results provided the first clear evidence for magnetic sensitivity in a crustacean. Arendse and Kruyswijk (1981) also reported that amphipods in the process of jumping aligned themselves with the field, whereas crawling amphipods did not. This provided a possible

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Fig. 12.2. Orientation of the marine isopod Idotea baltica basteri tested in an arena under two different magnetic fields. (A) Isopods tested in a natural magnetic field oriented along a magnetic axis that coincided with the seaward-landward axis in their natural environment. (B) Isopods tested in an artificial magnetic field in which magnetic north (MN) was rotated counterclockwise by 90 degrees showed a corresponding shift in orientation. Each circle or half circle on the diagram reflects the orientation of one isopod. Full circles indicate the direction of individuals that had unimodal orientation (i.e., moved consistently in one direction). Half circles indicate individuals with bimodal orientation (i.e., they moved in one direction in approximately half of the trials and the opposite direction in the other half). The double-headed arrows in the center of each circle indicate the axis along which each group of isopods moved. Diagram is modified from Ugolini and Pezzani (1995), with permission from Elsevier.

explanation for why earlier studies, which did not distinguish between crawling and jumping animals, might have failed to provide evidence for magnetic orientation. A subsequent study in Italy, however, again failed to find evidence for magnetic orientation (Scapini and Quochi 1992). It is possible that the different outcomes reflect population differences between amphipods in Italy and The Netherlands, or perhaps a difference in the importance of various local cues present at the specific beaches where the different studies were done (Wiltschko and Wiltschko 1995a). In an additional study carried out in Italy, Ugolini (1994) reported that altering the ambient field affected the orientation of Talitrus, but the quadrimodal orientation he observed differed from the bimodal orientation observed previously in The Netherlands. Similar experiments with the African amphipod Talorchestia martensii demonstrated that individuals oriented along the Y-axis of their home beach in complete darkness under the natural magnetic field, but oriented randomly when the magnetic field was canceled (Ugolini and Pardi 1992). Subsequent experiments designed to investigate interactions between the magnetic compass and the sun compass revealed that, when amphipods were tested with a view of the sun but in a magnetic field rotated 90 degrees clockwise, orientation was quadrimodal. Approximately half of the animals oriented along an axis coinciding with the real land-sea axis, whereas the other half oriented along the magnetic axis that had formerly coincided with the land-sea axis. These results are consistent with the hypothesis that the magnetic compass and sun compass are both used in Y-axis orientation, and that different individuals preferentially weigh one or the other if the cues are placed in conflict, a situation that never arises in nature. Additional experiments have shown that, when solar cues are absent, these amphipods use the geomagnetic field as the primary orientation cue (Ugolini et al. 1999, Ugolini 2001, 2002). Ugolini and Pezzani (1995) reported similar findings in the marine isopod Idotea baltica. Like the amphipods, Idotea oriented along the land-sea axis when tested in the natural magnetic field. When the ambient field was rotated to a new position, orientation shifted accordingly (Fig. 12.2). When the horizontal component of the field was canceled, orientation became random.



The Geomagnetic Sense of Crustaceans and Its Use in Orientation and Navigation

The isopods were also able to learn to orient in magnetic directions corresponding to directions that take them up or down slope, a response that would presumably help guide Y-axis orientation under natural conditions. Isopods exposed to a slope aligned perpendicular to the actual land-sea axis quickly learned to move along the magnetic axis corresponding to the new up-down slope direction, a directional preference expressed even when testing was done on a horizontal surface. These results imply that isopods can determine the land-sea axis in their environment using slope, and can then use a magnetic compass to move in appropriate directions during Y-axis orientation.

A MAGNETIC COMPASS IN SPINY LOBSTERS A magnetic compass sense has also been discovered in the Caribbean spiny lobster P. argus. In some geographic areas, this species undergoes an annual mass migration in which thousands of lobsters vacate shallow, inshore areas and crawl seaward in single-file, head-to-tail processions (reviewed by Kanciruk and Herrnkind 1978, Herrnkind 1980). Lines of spiny lobsters within the same geographical area follow nearly identical compass bearings (Herrnkind et al. 1973). Field and laboratory experiments have demonstrated that spiny lobsters can detect wave surge (the horizontal movement of water near the ocean floor) and use it as a directional cue (Walton and Herrnkind 1977, Nevitt et  al. 1995). Migratory orientation persists, however, in areas where hydrodynamic cues are disrupted or absent, and when visual cues are obscured by turbid water or darkness (Herrnkind 1970, Herrnkind and McLean 1971). In an initial attempt to determine whether spiny lobsters orient magnetically, Walton and Herrnkind (1977) captured lobsters in nonmigratory condition, covered their eyes, and displaced them to a new location. The orientation of ten lobsters carrying magnets was compared to the orientation of the same individuals carrying nonmagnetic solder wire as a control. Under these conditions, lobsters oriented into the prevailing wave surge regardless of whether magnets were attached. Although the results provided no evidence that spiny lobsters orient magnetically, the possibility that they use magnetic cues under different circumstances (e.g., in the absence of wave surge) could not be ruled out. In a subsequent laboratory experiment, an attempt was made to condition spiny lobsters to orient toward specific magnetic directions to receive food rewards (Lohmann 1985). Several lobsters were trained and tested in a rotatable circular orientation apparatus in which they could enter tunnels aligned with different compass directions. Two lobsters that had received positive reinforcement for orienting along a north-south magnetic axis were tested in the Earth’s field and in a field in which magnetic north was rotated 60 degrees clockwise. In each case, lobsters entered the tunnels aligned with magnetic north-south more often than expected by chance, providing initial evidence for magnetic sensitivity. Direct evidence for a magnetic compass in spiny lobsters was subsequently obtained in experiments carried out inside an underwater magnetic coil system located on a patch reef in the Florida Keys (Lohmann et al. 1995; Fig. 12.3). Lobsters were captured on the reef, and their eyestalks were covered with rubber eye caps to eliminate visual cues. Each animal was then tethered on a flat Plexiglas surface inside the coil. As the lobster walked, it was held in place by the tether, but its feet slipped across the surface. Most lobsters tethered in this way established and maintained consistent headings toward specific directions. After a lobster had established a consistent heading, it was exposed to either a reversal of the horizontal component of the Earth’s field, or to no change in the ambient field (controls). Those animals subjected to the field reversal deviated significantly from their initial courses, whereas

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Fig. 12.3. Underwater magnetic coil system and tethering procedure used to demonstrate the existence of a magnetic compass in spiny lobsters (from Lohmann et al. 1995, with permission from The Company of Biologists). See text for details.

control lobsters did not (Fig. 12.4). These results demonstrated that spiny lobsters have a magnetic compass sense and can use it to maintain headings in their natural habitat. Two functionally different types of magnetic compasses have been reported in animals. Polarity compasses, which are present in salmon (Quinn et al. 1981) and mole rats (Marhold et al. 1997), determine north using the polarity of the horizontal field component. By contrast, the inclination compasses of birds (Wiltschko and Wiltschko 1972) and sea turtles (Light et al. 1993, Goff et  al. 1998)  evidently do not detect the polarity of the field (i.e., north vs. south). Instead, they define “poleward” as the direction along the Earth’s surface in which the angle formed between the magnetic field vector and the gravity vector is smallest. Some salamanders have both types of compasses and use each in different behavioral tasks (Phillips 1986). To determine whether spiny lobsters have a polarity or inclination compass, Lohmann et al. (1995) exposed one group of lobsters tested in the underwater coil to a field with the vertical component of the field inverted, a treatment that does not affect animals with a polarity compass but that elicits a reversal of orientation direction in animals with an inclination compass.



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Fig. 12.4. Maximum deviations of spiny lobsters from initial paths established in the Earth’s magnetic field. The direction that each lobster walked initially was normalized to 0 degrees. (A) In trials in which the field was not changed, lobsters seldom deviated from the initial heading by more than 90 degrees. (B) In trials in which the horizontal field direction was reversed, lobsters had an average maximum deviation of 180 degrees. The arrows in the center of each circle indicate significant mean angles; the length of the arrow is proportional to the mean vector length r, with r = 1 represented by the edge of the circle. Dotted lines represent 95% confidence intervals for the mean angles. Diagram is modified from Lohmann et al. 1995, with permission from The Company of Biologists.

The orientation of the lobsters did not change, implying that they have a polarity compass functionally similar to that of salmon and mole rats. Although spiny lobsters have a magnetic compass, whether it is used to guide the autumn migration is not known. The hypothesis appears plausible but has not yet been tested experimentally, inasmuch as all studies on magnetic orientation have been carried out with lobsters that are not in migratory condition.

HOMING IN SPINY LOBSTERS The Caribbean spiny lobster is capable of at least some navigational tasks that cannot be explained by a magnetic compass alone. Working in Bermuda, Creaser and Travis (1950) captured and marked a number of spiny lobsters before displacing them to various locations. A surprising number (about 20%) were recaptured after displacement, including some that were released in deep water (1500 m) and had to travel in excess of 8 km to return to the capture sites. These findings led Creaser and Travis (1950) to conclude that these lobsters possess “a remarkable homing instinct,” the basis of which was unknown at the time. Subsequent studies revealed that the ability to return reliably to a home area appears to be a natural component of the behavior of this species. Juvenile and adult spiny lobsters spend daylight hours hidden inside coral reef crevices or holes, emerging at night to forage over a considerable area before returning in darkness to the same den or to one of several others nearby (Herrnkind et al. 1975, Herrnkind and Redig 1975).

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TRUE NAVIGATION IN SPINY LOBSTERS In the animal navigation literature, an animal is said to be capable of true navigation if, after displacement to a location where it has never been, it can determine its position relative to a goal without relying on familiar surroundings, cues that emanate from the destination, or information collected during the outward journey. In an experiment designed to determine whether spiny lobsters are capable of true navigation, lobsters in the Florida Keys, United States, were captured and transported by indirect, circuitous routes to test sites 12–37 km away from the capture site (Boles and Lohmann 2003). The animals were transported inside closed, opaque plastic containers partly filled with sea water, preventing access to visual cues and ensuring that they could not access chemical cues along the way. In some experiments, lobsters were also subjected to strong, varying magnetic fields during transport, to ensure that they could not use their magnetic compass to monitor the outward trip. At the test site, lobsters were permitted to sit overnight in the undisturbed magnetic field of the Earth before their orientation was tested the following morning. During tests, the lobsters were tethered so that they could walk in place inside a water-filled orientation arena. To ensure that the lobsters had never visited the test sites previously, the arena was placed on land. To prevent access to visual cues, the eyestalks of each animal were covered with rubber eye caps (Lohmann et al. 1995). Remarkably, lobsters tested in this way oriented in directions that, on average, coincided with paths back toward the site of capture (Fig. 12.5). These findings are consistent with the earlier homing studies of Creaser and Travis (1950) and imply that spiny lobsters are somehow able to determine their position relative to the capture site. Moreover, given that the animals were deprived of all known sources of positional information during transport, spiny lobsters appear to assess their geographic location based on information present at the test site (Boles and Lohmann 2003).

MAGNETIC MAPS IN SPINY LOBSTERS How spiny lobsters determine their position after being displaced to locations where they have never been was not immediately apparent. However, several features of the Earth’s field vary across the globe in such a way that they might, in principle, be used in position finding (Fig.  12.1). For example, at each geographic location, the magnetic field lines intersect the Earth’s surface at a specific angle of inclination. At the magnetic equator, the field lines are parallel to the Earth’s surface, and the inclination angle is said to be 0. The field lines become progressively steeper as one moves toward the magnetic poles; at the poles themselves, the field lines are perpendicular to the Earth’s surface. Thus, inclination angle varies predictably with latitude, and an animal able to detect this field element may be able to determine if it is north or south of a particular area. Sea turtles are known to be capable of this (Lohmann and Lohmann 1994). In addition to inclination angle, at least three other magnetic field elements vary across the Earth’s surface in ways that make them suitable for a position-finding sense (Skiles 1985, Lohmann et al. 1999, Putman et al. 2011). These include: (1) the intensity (strength) of the total field; (2) the intensity of the horizontal field; and (3) the intensity of the vertical field. To determine whether spiny lobsters possess “magnetic maps” that can be used to assess their position relative to their home areas, lobsters captured in the middle Florida Keys were placed into a circular pool of water so that their orientation could be monitored as described



The Geomagnetic Sense of Crustaceans and Its Use in Orientation and Navigation Moved from CS1 to TS1 0° 30°N 26°N 270°

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Fig. 12.5. Homing in displaced spiny lobsters. Spiny lobsters were transported by boat from two capture sites (CS1, CS2) via circuitous routes to one of two test sites (TS1, TS2). In the orientation diagrams, each small symbol represents the mean angle of a single spiny lobster. Squares indicate spiny lobsters captured at CS1, whereas circles indicate spiny lobsters captured at CS2. The arrow in the center of each orientation diagram indicates the mean angle of each group and the dotted lines represent the 95% confidence interval for the mean. The arrow outside each orientation diagram indicates the direction from the test site to the capture site. In each case, the mean angle of orientation coincided closely with the direction toward the capture site. Modified from Boles and Lohmann (2003), with permission from the Nature Publishing Group.

previously. This time, however, a large magnetic coil system was constructed around the pool so that lobsters could be tested in specific fields replicating those that exist in particular geographic areas (Boles and Lohmann 2003). Lobsters tested in a field that exists north of the capture site oriented southward, whereas those tested in a field like one that exists south of the capture site oriented northward (Fig. 12.6). These findings indicate that lobsters exploit magnetic information as a component of a classical navigational map which facilitates navigation to specific geographic locations. A lobster’s ability to navigate back to its home area appears to be based at least partly on the animal’s experience and learned understanding of how the Earth’s field varies in the geographic region where it lives (Lohmann and Lohmann 2006).

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Fig. 12.6. Evidence for a magnetic map in spiny lobsters. The diagram shows orientation of spiny lobsters tested in magnetic fields replicating those that exist at two different geographic locations (marked by stars on the map). Spiny lobsters were captured at Grassy Key (CS2 on Fig. 12.5), transported to the testing site (TS1 on Fig. 12.1), and held overnight in the local magnetic field before being tested in the morning. Spiny lobsters tested in a field like one that exists north of the test site walked southward, whereas those tested in a field like one that exists south of the test site walked northward. The open triangle outside each orientation diagram indicates the actual direction to the capture site from the test site. In each case, spiny lobsters responded as if they had been displaced to the locations marked by the stars rather than by orienting in the direction that was actually toward the capture site. Modified from Boles and Lohmann (2003), with permission from the Nature Publishing Group.

MECHANISM OF MAGNETIC FIELD DETECTION The mechanism or mechanisms underlying magnetic field detection have not been clearly established in any crustacean. Indeed, a full understanding of how any animal perceives magnetic fields has not yet been attained (Wiltschko and Wiltschko 2005, Johnsen and Lohmann 2005, 2008, Lohmann 2010). In recent years, most discussion of possible mechanisms underlying magnetoreception has focused on three main ideas: electromagnetic induction, magnetite, and chemical magnetoreception. We will consider these in turn, with special reference to the relevance of each to crustaceans. Electromagnetic Induction When an object composed of an electrically conductive material moves through a magnetic field in any direction other than parallel to the field lines, positively and negatively charged particles



The Geomagnetic Sense of Crustaceans and Its Use in Orientation and Navigation

migrate to opposite sides of the object, resulting in a constant voltage that depends on the speed and direction of the object’s motion relative to the magnetic field (Purcell 1985, Johnsen and Lohmann 2005, 2008). If the object is immersed in sea water (or any other conductive medium) that is stationary relative to the field, an electric circuit is formed and current flows through the medium and object. This principle of electromagnetic induction might explain how elasmobranch fish (sharks, skates, and rays) perceive magnetism (Kalmijn 1974, 1984). Because the bodies of these animals are conductive and the fish have highly sensitive electroreceptors, elasmobranchs might detect the voltage drop of the induced current that arises as they swim through Earth’s field (Lohmann and Johnsen 2000, Johnsen and Lohmann 2008). However, whether these fish actually perceive magnetic fields in this way is not known. In contrast to the situation in elasmobranchs, structures that serve as electroreceptors have never been identified in crustaceans. Recent reports, however, have provided behavioral evidence that crayfish may nevertheless perceive weak electric fields, an ability hypothesized to function in helping the animals detect hidden prey (Patullo and Macmillan 2007, 2010). If electroreceptors are confirmed to exist in crustaceans, then a magnetoreception mechanism based on electromagnetic induction is hypothetically possible. At present, however, the reported level of sensitivity to electrical stimuli by crayfish appears insufficient for crustaceans to exploit this mechanism in sensing the Earth’s magnetic field (Patullo and Macmillan 2010), or perhaps even in sensing prey (Steullet et al. 2007). Magnetite Some bacteria and unicellular algae orient their movements along magnetic field lines (Bazylinski and Frankel 2004). The discovery that crystals of the magnetic minerals magnetite (Fe3O4) and greigite (Fe3S 4) underlie this ability has inspired searches for similar minerals in diverse animals. Magnetite was subsequently detected in birds, salmon, sea turtles, and a number of other animals that are known to orient to the Earth’s magnetic field (Kirschvink et al. 1985). Most magnetite isolated from animals has been in the form of single-domain magnetite crystals similar to those found in magnetotactic bacteria (Johnsen and Lohmann 2005, 2008). Single-domain magnetite crystals are minute (about 50 nm in diameter), permanently magnetized magnets that twist into alignment with the Earth’s magnetic field if allowed to rotate freely. In principle, such crystals might transduce geomagnetic field information to the nervous system in several different ways. One possibility is that magnetite crystals exert torque or pressure on secondary receptors (such as stretch receptors, hair cells, or mechanoreceptors) as the particles attempt to align with the geomagnetic field. Alternatively, the rotation of intracellular magnetite crystals might open ion channels directly if, for example, cytoskeletal filaments connect the crystals to the channels. Additional ways that magnetite might interact with the nervous system have also been proposed (see Kirschvink et al. 2001, Johnsen and Lohmann 2005, Walker 2008 for more examples). Magnetic material that might be involved in magnetoreception has been detected in the Caribbean spiny lobster P.  argus (Lohmann 1984). The material is concentrated primarily in the cephalothorax, especially in tissue associated with the fused thoracic ganglia. Although direct evidence that magnetite underlies magnetoreception in spiny lobsters has not been obtained, findings consistent with this hypothesis have been acquired through pulse magnetization experiments (Cain 2001). A strong magnetic field of brief duration can be used to alter the direction of magnetization in magnetite particles. Pulse magnetization might, therefore, alter magnetite-based magnetoreceptors and change the behavior of animals that use such receptors

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Kenneth J. Lohmann and David A. Ernst to derive directional or positional information from the Earth’s field. In studies with several animals, including spiny lobsters (Cain 2001), the application of strong magnetic pulses either randomized the preferred orientation direction or else deflected it relative to controls (Wiltschko and Wiltschko 1995b, Beason et al. 1997, Irwin and Lohmann 2005). These results have generally been interpreted as evidence for magnetite-based magnetoreceptors, although other explanations cannot be ruled out with certainty (Johnsen and Lohmann 2005). In principle, strong magnetic pulses might alter magnetite-based receptors that are part of a compass sense, a map sense, or both. Additional research will be needed to determine whether magnetoreception in spiny lobsters is mediated by magnetite-based receptors. Chemical Magnetoreception A third hypothesis proposes that magnetoreception involves unusual biochemical reactions that are influenced by the Earth’s magnetic field. Because these reactions involve pairs of free radicals as fleeting intermediates, this idea is also known as the radical pairs hypothesis. The details of this proposed mechanism are complex and have been described elsewhere (Johnsen and Lohmann 2008, Rodgers and Hore 2009, Ritz et al. 2010). If chemical magnetoreception occurs, then it may be associated with the visual system (Liedvogel and Mouritsen 2010, Ritz et al. 2010). Many of the best-known radical-pair reactions begin with electron transfers that are induced by the absorption of light (Johnsen and Lohmann 2008, Ritz et al. 2010). This has led to the suggestion that chemical magnetoreceptors might also be photoreceptors. The possible link to photoexcitation has also led to interest in blue-light-sensitive photoreceptive proteins known as cryptochromes (Liedvogel and Mouritsen 2010). Cryptochromes are attractive candidates for magnetoreceptors because they exist in diverse animals and have a chromophore that forms radical pairs after photoexcitation (Johnsen and Lohmann 2005). Some evidence consistent with the hypothesis that cryptochromes function in magnetoreception has been obtained in migratory birds (Rodgers and Hore 2009, Ritz et al. 2010). The strongest evidence for cryptochrome involvement, however, comes from experiments with the fruit fly Drosophila, in which flies were trained to enter one arm of a simple maze on the basis of magnetic-field conditions. Mutant flies lacking genes for cryptochrome were unable to perform this task, but magnetic sensitivity was restored when cryptochrome genes were inserted into the flies (Gegear et al. 2008). If animals perceive magnetic fields using chemical reactions that occur within the visual system, then it is possible that they see, superimposed on their visual field, an additional signal consisting of a pattern of lights or colors, which changes depending on the magnetic direction that the animal faces (Lohmann 2010). Some indirect evidence exists to support this idea. For example, birds failed to orient magnetically when the right eye (which is known to be dominant in tasks involving object perception) was covered with a frosted foil that blurred vision, a result consistent with the hypothesis that interactions exist between processing visual patterns and detecting magnetic directions (Stapput et al. 2010). In crustaceans, however, no studies have been carried out investigating the possibility of chemical magnetoreception.

FUTURE DIRECTIONS The study of magnetoreception in crustaceans is still in its infancy. At present, studies have been conducted with only a few species, but the existence of magnetic sensitivity in several



The Geomagnetic Sense of Crustaceans and Its Use in Orientation and Navigation

different groups (amphipods, isopods, and lobsters) suggests that the ability to perceive magnetic fields might be widespread among crustaceans, and perhaps among arthropods more generally (e.g., Arendse 1978, Baker 1987, Phillips and Sayeed 1993, Gegear et al. 2008). Investigations with diverse crustaceans appear likely to expand the list of species known to perceive magnetic fields. The Caribbean spiny lobster P.  argus is the most thoroughly studied crustacean in terms of magnetoreception, yet numerous questions remain about exactly how this species exploits geomagnetic information. For example, although the lobsters’ “magnetic map” is thought to rely on magnetic field parameters such as intensity and field inclination (Boles and Lohmann 2003), the precise way in which the map is organized is not understood (Lohmann and Lohmann 2006, Lohmann et al. 2007). Similarly, the capabilities and limitations of the map have not yet been investigated, nor is it known how the magnetic compass and map function during the seasonal migrations of these lobsters. Finally, almost nothing is known about the neural mechanisms that underlie magnetic field detection in crustaceans. Although the existence of magnetic material in spiny lobsters (Lohmann 1984) is consistent with the hypothesis that magnetite serves as the physical basis for the magnetic sense, definitive evidence for magnetite-based magnetoreception is lacking. Clearly, research on a number of different fronts appears likely to result in major advances.

SUMMARY AND CONCLUSIONS Representatives of several crustacean groups, including amphipods, isopods, and spiny lobsters, have magnetic compasses that enable them to orient movements relative to the Earth’s magnetic field. In addition, the Caribbean spiny lobster has a magnetic map sense, which allows it to use positional information in the Earth’s field to navigate toward its home area. Little is known about the mechanism(s) that underlie magnetic field detection in crustaceans. Evidence consistent with magnetite-based receptors has been obtained in the spiny lobster, but the possibility that additional or alternative mechanisms exist in this and other species cannot be excluded. Studies on magnetoreception in crustaceans are still at a very early stage. Given how few species and groups have been investigated so far, it appears possible that magnetoreception is a widespread sensory ability among crustaceans. Future studies in this area are likely to be rewarding.

REFERENCES Adamczewska, A.M., and S. Morris. 1998. Strategies for migration in the terrestrial Christmas Island red crab Gecarcoidea natalis: intermittent versus continuous locomotion. Journal of Experimental Biology 201:3221–3231. Arendse, M.C. 1978. Magnetic field detection is distinct from light detection in the invertebrates Tenebrio and Talitrus. Nature 274:358–362. Arendse, M.C. 1980. Non-visual orientation in the sandhopper Talitrus saltator (Mont.). Netherlands Journal of Zoology 30:535–554. Arendse, M.C., and C.J. Kruyswijk. 1981. Orientation of Talitrus saltator to magnetic fields. Netherlands Journal of Sea Research 15:23–32. Baker, R.R. 1987. Integrated use of moon and magnetic compass by the heart-and-dart moth, Agrotis exclamationis. Animal Behaviour 35:691–704.

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Kenneth J. Lohmann and David A. Ernst Bazylinski, D.A., and R.B. Frankel. 2004. Magnetosome formation in prokaryotes. Nature Reviews Microbiology 2:217–230. Beason, R.C., R. Wiltschko, and W. Wiltschko. 1997. Pigeon homing: effects of magnetic pulses on initial orientation. Auk 114:405–415. Boles, L.C., and K.J. Lohmann. 2003. True navigation and magnetic maps in spiny lobsters. Nature 421:60–63. Cain, S.D. 2001. The neuroethology of magnetic orientation behavior in two invertebrate animals. PhD dissertation, University of North Carolina, Chapel Hill, North Carolina, USA. Creaser, E.P., and D. Travis. 1950. Evidence of a homing instinct in the Bermuda spiny lobster. Science 11:169–170. Cubit, J. 1969. Behavior and physical factors causing migration and aggregation of the sand crab Emerita analoga (Stimpson). Ecology 50:118–123. Ercolini, A., and F. Scapini. 1972. On the non-visual orientation of littoral amphipods. Monitore Zoologico Italiano (N.S.) 6:75–84. Forward Jr., R.B., H. Diaz, and J.H. Cohen. 2005. The tidal rhythm of the mole crab Emerita talpoida. Journal of the Marine Biological Association of the United Kingdom 85:895–901. Gegear, R.J., A. Casselman, S. Waddell, and S.M. Reppert. 2008. Cryptochrome mediates light-dependent magnetosensitivity in Drosophila. Nature 454:1014–1018. Goff, M., M. Salmon, and K.J. Lohmann. 1998. Hatchling sea turtles use surface waves to establish a magnetic compass direction. Animal Behaviour 55:69–77. Herrnkind, W.F. 1970. Migration of the spiny lobster. Natural History 79:36–43. Herrnkind, W.F. 1980. Spiny lobsters: patterns of movement. Pages 349–407 in J.S. Cobb and B.F. Phillips, editors. Biology and management of lobsters, Vol. 1. Physiology and behavior. Academic Press, New York. Herrnkind, W.F., and P. Kanciruk. 1978. Mass migration of spiny lobster, Panulirus argus (Crustacea: Palinuridae): synopsis and orientation. Pages 430–439 in K. Schmidt-Koenig and W.T. Keeton, editors. Animal migration, navigation, and homing. Springer-Verlag, New York. Herrnkind, W.F., and R. McLean. 1971. Field studies of homing, mass emigration and orientation in the spiny lobster, Panulirus argus. Annals of the New York Academy of Sciences 188:359–377. Herrnkind, W.F., and M.X. Redig. 1975. Preliminary study of establishment of den residency by spiny lobster, Panulirus argus, at Grand Bahama Island. Hydro-Lab Journal 3:96–101. Herrnkind, W.F., P. Kanciruk, J. Halusky, and R. McLean. 1973. Descriptive characterization of mass autumnal migrations of spiny lobster, Panulirus argus. Proceedings of the Gulf and Caribbean Fisheries Institute 25:78–98. Herrnkind, W.F., J.A. Van DerWalker, and L. Barr. 1975. Population dynamics, ecology and behavior of spiny lobsters, Panulirus argus, of St. John, U.S.V.I. IV. Habitation, patterns of movement and general behavior. Science Bulletin of the Museum of Natural History, Los Angeles County 20:31–45. Hughes, D. 1966. Behavioural and ecological investigations of the crab Ocypode ceratophthalmus (Crustacea: Ocypodidae). Journal of Zoology 150:129–143. Irwin, W.P., and K.J. Lohmann. 2005. Disruption of magnetic orientation in hatchling loggerhead sea turtles by pulsed magnetic fields. Journal of Comparative Physiology A 191:475–480. Johnsen, S., and K.J. Lohmann. 2005. The physics and neurobiology of magnetoreception. Nature Reviews Neuroscience 6:703–712. Johnsen, S., and K.J. Lohmann. 2008. Magnetoreception in animals. Physics Today 61:29–35. Kalmijn, A.J. 1974. The detection of electric fields from inanimate and animate sources other than electric organs. Pages 147–200 in A. Fessard, editor. Handbook of sensory physiology, Vol. III/3. Springer, New York. Kalmijn, A.J. 1984. Theory of electromagnetic orientation: a further analysis. Pages 525–560 in L. Bolis, R.D. Keynes, and H.P. Maddrell, editors. Comparative physiology of sensory systems. Cambridge University Press, Cambridge, UK. Kanciruk, P., and W.F. Herrnkind. 1978. Mass migration of spiny lobster, Panulirus argus (Crustacea: Palinuridae): behavior and environmental correlates. Bulletin of Marine Science 28:601–623.



The Geomagnetic Sense of Crustaceans and Its Use in Orientation and Navigation

Kirschvink, J.L., D.S. Jones, and B.J. MacFadden. 1985. Magnetite biomineralization and magnetoreception in organisms. Plenum Press, New York. Kirschvink, J.L., M.M. Walker, and C.E. Diebel. 2001. Magnetite-based magnetoreception. Current Opinion in Neurobiology 11:462–467. Liedvogel, M., and H. Mouritsen. 2010. Cryptochromes—a potential magnetoreceptor: what do we know and what do we want to know? Journal of Royal Society Interface 7:S147–S162. Light, P., M. Salmon, and K.J. Lohmann. 1993. Geomagnetic orientation of loggerhead sea-turtles— evidence for an inclination compass. Journal of Experimental Biology 182:1–9. Lohmann, K.J. 1984. Magnetic remanence in the western Atlantic spiny lobster, Panulirus argus. Journal of Experimental Biology 113:29–41. Lohmann, K.J. 1985. Geomagnetic field detection by the western Atlantic spiny lobster, Panulirus argus. Marine and Freshwater Behaviour and Physiology 12:1–17. Lohmann, K.J. 2010. Magnetic-field perception: News and Views Q&A. Nature 464:1140–1142. Lohmann, K.J., and S. Johnsen. 2000. The neurobiology of magnetoreception in vertebrate animals. Trends in Neurosciences 23:153–159. Lohmann, K.J., and C.M.F. Lohmann. 1994. Detection of magnetic inclination angle by sea turtles: a possible mechanism for determining latitude. Journal of Experimental Biology 194:23–32. Lohmann, K.J., and C.M.F. Lohmann. 2006. Sea turtles, lobsters, and oceanic magnetic maps. Marine and Freshwater Behaviour and Physiology 39:49–64. Lohmann, K.J., N.D. Pentcheff, G.A. Nevitt, G. Stetten, R.K. Zimmer-Faust, H.E. Jarrard, and L.C. Boles. 1995. Magnetic orientation of spiny lobsters in the ocean: experiments with undersea coil systems. Journal of Experimental Biology 198:2041–2048. Lohmann, K.J., J.T. Hester, and C.M.F. Lohmann. 1999. Long-distance navigation in sea turtles. Ethology Ecology and Evolution 11:1–23. Lohmann, K.J., C.M.F. Lohmann, and N.F. Putman. 2007. Magnetic maps in animals: nature’s GPS. Journal of Experimental Biology 210:3697–3705. Lohmann, K.J., C.M.F. Lohmann, and C.S. Endres. 2008. The sensory ecology of ocean navigation. Journal of Experimental Biology 211:1719–1728. Lohmann, K.J., N.F. Putman, and C.M.F. Lohmann. 2012. The magnetic map of hatchling loggerhead sea turtles. Current Opinion in Neurobiology 22:336–342. Marhold, S., W. Wiltschko, and H. Burda. 1997. A magnetic polarity compass for direction finding in a subterranean mammal. Naturwissenschaften 84:421–423. Nevitt, G.A., N.D. Pentcheff, K.J. Lohmann, and R.K. Zimmer-Faust. 1995. Hydrodynamic orientation by spiny lobsters: field manipulations in a patch reef environment. Journal of Experimental Biology 198:2049–2054. Pardi, L., and A. Ercolini. 1986. Zonal recovery mechanisms in talitrid crustaceans. Italian Journal of Zoology 53:139–160. Pardi, L., and F. Scapini. 1983. Inheritance of solar direction finding in sandhoppers—mass-crossing experiments. Journal of Comparative Physiology A 151:435–440. Patullo, B.W., and D.L. Macmillan. 2007. Crayfish respond to electrical fields. Current Biology 17:R83–R84. Patullo, B.W., and D.L. Macmillan. 2010. Making sense of electrical sense in crayfish. Journal of Experimental Biology 213:651–657. Phillips, J.B. 1986. Two magnetoreception pathways in a migratory salamander. Science 233:765–767. Phillips, J.B., and O. Sayeed. 1993. Wavelength-dependent effects of light on magnetic compass orientation in Drosophila melanogaster. Journal of Comparative Physiology A 172:303–308. Purcell, E.M. 1985. Electricity and magnetism: Berkeley physics course, Vol. 2. McGraw-Hill, New York. Putman, N.F., C.S. Endres, C.M.F. Lohmann, and K.J. Lohmann. 2011. Longitude perception and bicoordinate magnetic maps in sea turtles. Current Biology 21:463–466. Quinn, T.P., R.T. Merrill, and E.L. Brannon. 1981. Magnetic field detection in sockeye salmon. Journal of Experimental Zoology 217:137–142. Ritz, T., M. Ahmad, H. Mouritsen, R. Wiltschko, and W. Wiltschko. 2010. Photoreceptor-based magnetoreception: optimal design of receptor molecules, cells, and neuronal processing. Journal of the Royal Society Interface 7:S135–S146.

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Kenneth J. Lohmann and David A. Ernst Rodgers, C.T., and P.J. Hore. 2009. Chemical magnetoreception in birds: the radical pair mechanism. Proceedings of the National Academy of Sciences of the United States of America 106:353–360. Scapini, F. 2006. Keynote papers on sandhopper orientation and navigation. Marine and Freshwater Behaviour and Physiology 39:73–85. Scapini, F., and G. Quochi. 1992. Orientation in sandhoppers from Italian populations: have they magnetic orientation ability? Monitore Zoologico Italiano (N. S.) 59:437–442. Stapput, K., O. Gunturkun, K.P. Hoffmann, R. Wiltschko, and W. Wiltschko. 2010. Magnetoreception of directional information in birds requires nondegraded vision. Current Biology 20:1259–1262. Skiles, D.D. 1985. The geomagnetic field: its nature, history and biological relevance. Pages 43–102 in J.L. Kirschvink, D.S. Jones, and B.J. MacFadden, editors. Magnetite biomineralization and magnetoreception in organisms. Plenum Press, New York. Steullet, P., D.H. Edwards, and C.D. Derby. 2007. An electric sense in crayfish? Biological Bulletin 213:16–20. Tankersley, R.A., M.G. Wieber, M.A. Sigala, and K.A. Kachurak. 1998. Migratory behavior of ovigerous blue crabs Callinectes sapidus: evidence for selective tidal-stream transport. Biological Bulletin 195:168–173. Ugolini, A. 1994. Magnetic orientation in and near the sea: Idotea baltica, Talitrus saltator and Talorchestia martensii. Page 10 in R. Williamson, editor. Orientation and migration at sea. Marine Biological Association and Society for Experimental Biology, Plymouth, United Kingdom. Ugolini, A. 2001. Relationship between compass systems of orientation in equatorial sandhoppers. Animal Behaviour 62:193–199. Ugolini, A. 2002. The orientation of equatorial sandhoppers during the zenithal culmination of the sun. Ethology Ecology and Evolution 14:269–273. Ugolini, A., and L. Pardi. 1992. Equatorial sandhoppers do not have a good clock. Naturwissenschaften 79:279–281. Ugolini, A., and A. Pezzani. 1995. Magnetic compass and learning of the Y-axis (sea-land) direction in the marine isopod Idotea baltica basteri. Animal Behaviour 50:295–300. Ugolini, A., G. Beugnon, and M. Vannini. 1988. Orientation and escape reaction in Palaemonetes antennarius (Milne Edwards) (Natantia Palaemonidae). Monitore Zoologico Italiano (N.S.) 22:105–109. Ugolini, A., B. Vignali, and P. Posso. 1999. Talorchestia tricornuta Shoemaker (Amphipoda, Talitridae) from Sandy Shores of Gabon: compass mechanisms of orientation. Ethology 105:25–36. Van den Bercken, J., S. Broekhuizen, J. Ringelberg, and H.H.W. Velthuis. 1967. Non-visual orientation in Talitrus saltator. Experientia 23:44–45. Vannini, M., and S. Cannicci. 1995. Homing behavior and possible cognitive maps in crustacean decapods. Journal of Experimental Marine Biology and Ecology 193:67–91. Walker, M.M. 2008. A model for encoding of magnetic field intensity by magnetite-based magnetoreceptor cells. Journal of Theoretical Biology 250:85–91. Walton, A.S., and W.F. Herrnkind. 1977. Hydrodynamic orientation of the spiny lobster, Panulirus argus (Crustacea: Palinuridae): wave surge and unidirectional currents. Memorial University of Newfoundland Marine Sciences Research Laboratory, Technical Reports 20:184–211. Wiltschko, W., and R. Wiltschko. 1972. Magnetic compass of European robins. Science 176:62–64. Wiltschko, R., and W. Wiltschko. 1995a. Magnetic orientation in animals. Springer, Berlin, Germany. Wiltschko, W., and R. Wiltschko. 1995b. Migratory orientation of European robins is affected by the wavelength of light as well as by a magnetic pulse. Journal of Comparative Physiology A 177:363–369. Wiltschko, W., and R. Wiltschko. 2005. Magnetic orientation and magnetoreception in birds and other animals. Journal of Comparative Physiology A 191:675–693. Zeil, J. 1998. Homing in fiddler crabs (Uca lactea annulipes and Uca vomeris: Ocypodidae). Journal of Comparative Physiology A 183:367–377.

13 THE CRUSTACEAN STOMATOGASTRIC NERVOUS SYSTEM

Eve Marder, Marie L. Goeritz, Gabrielle J. Gutierrez, Albert Hamood, Ted Brookings, Jonathan Caplan, Sara Haddad, Tilman Kispersky, and Sonal Shruti

Abstract This chapter provides an overview of the organization and function of the crustacean stomatogastric nervous system (STNS). The STNS generates motor patterns that move the approximately 40 sets of striated muscles of the animal’s stomach. The stomatogastric ganglion (STG) contains about 30 neurons (numbers vary slightly across crustacean species), including a large number of motor neurons and several interneurons. Motor neurons are identified unambiguously according to the muscles they innervate, and this has allowed the establishment of a connectivity diagram for the STG that includes many chemical inhibitory and electrical synapses. The continuous triphasic pyloric rhythm has a characteristic period of about 1 sec, while the episodic gastric mill rhythm has a characteristic period of about 5–10 sec. Descending modulatory neurons are activated by sensory inputs to inf luence the pyloric rhythm and to activate gastric mill rhythms. The large number of neuromodulatory substances found in these descending inputs include conventional small molecule neurotransmitters, amines, neuropeptides, and gases. Biophysical and molecular studies have characterized many of the voltage-dependent currents expressed in identified STG neurons. The interaction of these currents and synaptic strengths for circuit dynamics has been studied by combining electrophysiological and modeling studies. STG neurons and circuits have also been useful for understanding long-term homeostatic regulation of neuronal excitability and the development of motor patterns.

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INTRODUCTION The crustacean STNS is one of the premier preparations used to understand how cellular and synaptic properties give rise to circuit dynamics and the generation of rhythmic motor patterns (Selverston and Moulins 1987, Harris-Warrick et al. 1992, Marder and Bucher 2007, Stein 2009). The utility of this preparation for the past 40 years demonstrates the prescience of the early workers (Maynard 1972, Maynard and Dando 1974, Mulloney and Selverston 1974a, 1974b, Hartline and Maynard 1975, Selverston et al. 1976) who recognized its extraordinary attributes for the analysis of a circuit. These attributes include (1) the neurons were individually identifiable, (2) intracellular recordings from the somata were sufficient to reveal synaptic interactions, and (3) the isolated preparation produced fictive motor patterns very similar to those seen in vivo (Rezer and Moulins 1983, Heinzel 1988a, 1988b, Clemens et al. 1998b). The utility of the STNS to understand a myriad of problems associated with circuit dynamics endures precisely because there have been so few other preparations with these same attributes. In this chapter, we will first present some of the basic features of the STNS and then briefly present some of the many important insights into how circuits work that have come from the study of this important preparation. Sadly, because of length limitations, this chapter cannot do justice to the almost 1,000 research papers and two books (Selverston and Moulins 1987, Harris-Warrick et al. 1992) published to date on the STNS, nor can we adequately place this work into the conceptual frameworks and scientific problems to which it has contributed. Nonetheless, we hope this chapter whets the reader’s appetite for more of the fascinating work done during the past 40 years that exploits the unique features of the STNS for fundamental studies of numerous issues in neuroscience and biology. We apologize to our colleagues, past and present, for omitting specific citations to many wonderful papers that have contributed much to what we know. As recent years have brought new methods of circuit analysis and new interest in determining “the connectome” in animals and humans, the lessons learned from the STNS become even more significant, as it is one of very few preparations for which we have had synaptic connectivity diagrams long enough to understand how difficult the next steps in analyzing circuit dynamics can be.

THE STOMATOGASTRIC NERVOUS SYSTEM Gross Anatomy The STNS controls the movements of the crustacean foregut (Maynard and Dando 1974) and consists of four ganglia, the paired commissural ganglia (CoGs), the oesophageal ganglion (OG) the stomatogastric ganglion (STG), and their connecting nerves. Figure 13.1A is a schematic diagram of the foregut, showing the oesophagus, cardiac sac, gastric mill, and pyloric regions of the stomach, and the position of the STG. Food enters the oesophagus and moves into the cardiac sac before being chewed by the lateral and medial teeth of the gastric mill (for details see Watling 2013). Food then passes into the pyloric region, where it is sieved and filtered (Maynard and Dando 1974). Associated with these anatomical regions of the foregut are four foregut rhythms, the oesophageal rhythm, cardiac sac rhythm, gastric mill rhythm, and pyloric rhythms. The oesophageal rhythm has a characteristic period of about 10 sec, but the mechanisms by which it is generated remain unknown. The cardiac sac rhythm is quite slow, with a period of about 30 sec (Dickinson and Marder 1989), and although its interactions with the pyloric and gastric mill



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rhythm are relatively well-characterized (Dickinson and Marder 1989, Dickinson et al. 1990), again the mechanisms underlying its generation remain largely unknown. In contrast, the gastric mill and pyloric rhythms have been extensively studied in a variety of crustacean species, and the remainder of this chapter will focus on these two motor patterns and their control by modulatory neurons with somata in the CoGs and OG. The crustacean stomach is a complex structure, with more than 40 pairs of striated muscles (Maynard and Dando 1974) (Fig. 13.1A) that move the teeth and the stomach ossicles. These muscles are innervated by excitatory motor neurons, most of which are found in the STG; neurons are identified on the basis of the muscles that they innervate. Figure 13.1B is a schematic of the crab STNS as it would be laid out in a saline-filled dish for electrophysiological recordings. This diagram shows the major nerves of the STNS, and the projection patterns of the STG motor neurons. Figure  13.1C shows example recordings of the STG motor patterns from the crab Cancer borealis, recorded with a combination of somatic intracellular recordings and extracellular recordings made from the motor nerves. The top three traces are simultaneous extracellular recordings from the nerves indicated. The fast triphasic pyloric rhythm is seen as the alternating bursts of the pyloric dilator (PD), lateral pyloric (LP), and pyloric (PY) neurons on the lateral ventricular nerve (lvn) trace. The medial ventricular nerve (mvn) trace shows activity of the inferior cardiac (IC) and ventricular dilator (VD) neurons, which multiplex the pyloric and gastric rhythms. These neurons are firing in long bursts (in gastric time), during which time they are also firing in phase with the fast pyloric rhythm. The dorsal gastric nerve (dgn) trace shows slow alternating bursts of the dorsal gastric (DG) and gastric mill (GM) neurons, members of the gastric mill circuit. The fourth trace is an intracellular recording of the lateral gastric (LG) neuron (its activity is also seen on the lvn trace). The LG neuron depolarizes and fires almost in phase with the GM neurons, and in alternation with the DG neuron. While the basic features of the pyloric and gastric mill rhythms are conserved across species, there are minor species differences in the phase relationships of the STG neurons during these rhythms. These species differences in STG motor patterns almost certainly reflect the fact that the shape and proportions of the stomach change to accommodate the different body plans of lobsters and crabs. The connectivity among the neurons of the STG in the lobster Panulirus interruptus was established by combining intracellular recordings and photoinactivations of specific neurons subsequent to f luorescent dye fills (Miller and Selverston 1979, Selverston and Miller 1980, Eisen and Marder 1982, Marder and Eisen 1984). Because the large number of electrical synapses create the potential for ambiguity in determining the exact pattern of connectivity, cell deletions were necessary (Eisen and Marder 1982) to disambiguate the connections when electrical synapses are present. The connectivity among pyloric and gastric mill neurons in other species remains less certain because of ambiguities caused by ubiquitous electrical synapses (Fig. 13.1D). Morphology of Stomatogastric Ganglion Neurons The morphology of the STG neurons has been studied at the level of the light microscope using intracellular dye fills (Fig. 13.2) (Bucher et al. 2007, Thuma et al. 2009) and ultrastructurally using electron microscopy (Maynard 1971, King 1976a, 1976b, Kilman and Marder 1996, Cabirol-Pol et al. 2002). The exact position of the identified neurons of the STG varies from animal to animal, although there are often preferred groupings of cells and preferred positions along the anterior-posterior axis of the ganglion (Wilensky et al. 2003, Bucher et al. 2007). The large (60–120 μm diameter) unipolar neurons of the STG send a primary neurite (15–30 μm diameter) into the center of the ganglion, the coarse neuropil, which is mostly devoid of synaptic

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Fig. 13.1. The stomatogastric nervous system. (A) Side view of a generic crab stomach showing the position of the stomach regions, ganglia, major nerves, and stomach muscles (redrawn from Maynard and Dando 1974, with permission from The Royal Society). (B) Schematic of the dissected stomatogastric nervous system as it might appear laid out flat for recording. Lowercase abbreviations are the nerve names, and the neurons whose action potentials can be recorded at various places are shown in capital letters in the boxes. Modified from an unpublished drawing by J.M. Weimann and H.-G. Heinzel. (C) Simultaneous recordings of the pyloric and gastric rhythms of the crab Cancer borealis. Top three traces are extracellular recordings made from the nerves at the positions shown in B.  The bottom trace is an intracellular recording from the LG neuron. Unpublished recordings of G. Gutierrez. (D) Connectivity diagram of the crab STG. Electrical synapses denoted by resistor symbols, chemical inhibitory synapses by filled circles (modified from Marder and Bucher 2007, with permission from Annual Reviews).



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contacts (Fig. 13.2). The primary neurite splits into multiple branches of vastly different diameters that ramify into a shell of fine processes (100 individuals per m 2. Because they are confined geographically to estuaries and bays, which are separated by several hundred or thousand kilometers of oceanic coast, populations in different regions are thought to be isolated from one another. Along its wide, but discontinuous, geographic distribution, the species experiences different tidal regimes (from few cm to 9 m), water salinities (from near 0 to 60 ppt), environmental structures (mud flat vs. vegetated areas), and predation risks (areas where aerial predators are abundant or scarce) (Luppi et al. 2013). Consequently, N. granulata has been an excellent model for studying intra- and interpopulation variability (reviewed in Spivak 2010). Neohelice is a robust midsized running crab, reaching up to 36 mm across the carapace, which digs semipermanent burrows and is active both in air and water. It is a highly visual animal that displays conspicuous visually guided behaviors similar to those described in fiddler crabs (Zeil and Hemmi 2006). Each of its two eyes, containing around 8,000 ommatidia, is mounted on a 5-mm-long eyestalk. As in fiddler crabs, the eyes of Neohelice possess a zone of higher acuity around the equator (Berón de Astrada et  al. 2011). The neuroanatomy and physiology of the visual nervous system of Neohelice is probably the best known in any crustacean and will be described later. Neohelice strongly relies on visual information. Among other things, the visual system allows the animal to detect and organize anticipatory responses to the attacks of its aerial predators (references in Luppi et al. 2013). These responses, however, need to be adaptive, that is, they must be sensitive to modification by information acquired through new experiences. Neohelice: Learning in the Laboratory Our studies in the laboratory have shown the ability of Neohelice to acquire and retain different types of memory. Paradigms that proved to successfully induce long-term memory (24 h or more) include: (1) habituation of the escape response to a visual danger stimulus (e.g., Brunner and Maldonado 1988, Lozada et  al. 1990, Tomsic and Maldonado 1990); (2)  habituation of exploratory activity to the contextual environment (Dimant and Maldonado 1992); (3) sensitization to electrical shocks or to visual moving stimuli (Rakitin et al. 1991, Aggio et al. 1996); (4) passive avoidance learning (Denti et al. 1988, Fernandez-Duque et al. 1992); (5) appetitive conditioning (Dimant and Maldonado 1992, Tomsic et al. 1996, Dimant et al. 2002); and (6) contextual learning (Tomsic et al. 1993, 1998, Pedreira et al. 1996). Behavioral studies using some of these paradigms were occasionally followed up by pharmacological studies (e.g., Maldonado

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Daniel Tomsic and Héctor Maldonado et al. 1989, Valeggia et al. 1989, Romano et al. 1990, Tomsic et al. 1991, Aggio et al. 1996). However, the paradigm that has been used the most in Neohelice as a model for studying the neurobiology of learning and memory with a multidisciplinary approach is “context-signal memory” (CSM), as explained below. Context-Signal Memory in Neohelice Neohelice is preyed on by gulls and other sea birds; hence, an object moving overhead elicits the animal’s escape. In our laboratory, a black rectangular screen moving horizontally overhead represents a visual danger stimulus (VDS) that evokes the crab’s escape. A typical trial with a VDS consists of two back-and-forth circular movements of the screen over 9 sec (Fig. 19.1). The escape response is recorded with the animal located inside an actometer, which consists of a bowl-shaped container with circular f lat f loor covered to a depth of 0.5 cm with water. The response is captured by a transducer device (the system was modified and upgraded several times over the last 20 years) and recorded with a computer (Fig. 19.1). The experimental room has 40 actometers, isolated from each other by partitions, which allows simultaneous recordings from 40 crabs. The basic experimental design used for assessing long-term memory in the crab includes a training session and a testing session typically separated by 24 h, and at least two groups of animals, one trained and one untrained. Pharmacological or other intervening experiments include the corresponding additional control and trained pairs of groups (e.g., control and trained, both injected with a certain drug). Each group usually includes 30–40 crabs. In the first session, the trained group receives the repetitive presentation of the VDS, while the control group just remains in the actometers without any phasic stimulation. At the end of this session, the animals are put in individual containers, where they remain until the testing session. In the second session, all animals are put again in the actometers to be tested with the VDS. Memory retention is said to occur when the trained group shows a level of responsiveness to the VDS that is statistically lower than that of the control group. A  single training session with 15 presentations of the VDS separated by 3 min (spaced training) invariably causes a reduction of the escape response that is retained for more than 1 day (Fig. 19.2B) (Tomsic et al. 1998, 2003, Hermitte et al. 1999, Pedreira and Maldonado 2003, Sztarker and Tomsic 2011). Although at the beginning we termed this phenomenon “habituation” (e.g., Lozada et  al. 1990, Romano et  al. 1991, Tomsic et  al. 1991, 1993, 1996), later investigations demonstrated that it is a more complex form of memory. In fact, the long-term modification of the escape response is exhibited only if the animal is tested in the same visual environment where it was trained. In other words, a change in the visual context between training and testing prevents the memory from being evoked (Fig. 19.3) (Tomsic et al. 1998, Hermitte et al. 1999, Sztarker and Tomsic 2011). Moreover, exposure to the context alone, prior to or following the training, the typical procedure that causes respectively latent inhibition or extinction, impairs the formation or the expression of the crab’s memory (Tomsic et al. 1998). These results led to the conclusion that the memory produced by spaced training is determined by an association between the VDS and the context (Tomsic et al. 1998, Sztarker and Tomsic 2011). For this reason, during the last decade, we have called this associative memory “context-signal memory” (CSM). Thus, CSM entails an association between two visual memories, a memory of the context (CM) and a memory of the signal (SM), each of which can be acquired independently (Tomsic et  al. 1993, 1998, Hermitte et  al. 1999). More recent investigations continue to support these conclusions (Pedreira et al. 2002, 2004, Pedreira and Maldonado 2003, Sztarker and Tomsic 2011).



Neurobiology of Learning and Memory of Crustaceans A

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Fig. 19.1. Experimental setups and recording procedures. (A) Behavioral experiments were conducted in an actometer that translated crab movement into voltage changes. (B)  For intracellular recording, the crab was held in an adjustable clamp. The eyestalks were cemented to the carapace, and a glass microelectrode was advanced through a small hole in the cuticle made at the tip of one eye. (C) Representative recording (in arbitrary units) of the escape response to a visual danger stimulus (VDS) during a trial (two screen motion cycles). (D) Representative recording of the response of a lobula giant (LG) neuron to the same stimulation. In C and D, the two topsides of the triangles below the traces represent the clockwise and counterclockwise movements of the stimulus contained in a cycle, and the base of the triangle represents 2.2 sec of record. From Sztarker and Tomsic (2008), with permission from Springer.

The Adaptive Value of the Context-Signal Memory Suppressing an escape response to a potentially dangerous stimulus for a long period of time can be highly risky. Then, why is Neohelice so prone to give up escaping to a VDS for such a long time? To answer this question, further information about the characteristics of learning responses to danger stimuli is needed. First, regarding a stimulus that signals danger in the wild, the greater the ambiguity of the signal, the greater the likelihood to stop responding to such stimulus. Thus, a high degree of behavioral change is expected when the stimulus almost invariably proves to be innocuous, in contrast to the lack of change expected when an unequivocal relationship links the stimulus and a subsequent damage. Second, the learning-induced change of response to VDS in Neohelice is stimulus specific (Lozada et al. 1990). In fact, animals tested with VDS that were different (or moved differently) than in the training escaped like untrained animals. In other words, crabs are capable of recognizing a visual motion stimulus

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Fig. 19.2. Short- and long-term memory of groups of crabs that received massed (A) or spaced (B) training. During the first day, animals in the trained groups (circles) received the repetitive presentation of a visual danger stimulus (VDS), while those of the control groups remained in the actometer without being stimulated. The following day, all animals were tested with the VDS. (A) Massed training consisted of 300 trials without intertrial intervals (ITI = 0 sec) (45-min session). Trained animals first displayed a conspicuous escape response that waned rapidly and deeply after a few trials. However, 24 h later no difference between trained and control (triangles) groups were observed. (B) Spaced training consisted of 15 trials with an ITI of 3 min. The training protocol lasted 45 min. The rate of reduction of the escape response during training was slower than with massed training. However, only 15 trials of spaced training resulted in a response reduction that was long-lasting, as disclosed 24 h later by a significant difference with respect to the control group. Graphs show means SEM. *p < 0.01. From Tomsic et al. (2009), with permission from Elsevier.

that proved to be innocuous from a slightly different one (Sztarker and Tomsic 2011). But are there innocuous visual motion stimuli in the natural environment of Neohelice that need to be ignored? The answer is yes. This species inhabits an upper intertidal zone densely vegetated by cord grass Spartina alterniflora or Spartina densiflora, both erect, tough, long-leaved grasses ranging from 0.3 to 2 m tall. Therefore, the high propensity of Neohelice to reduce its escape response to an iterated object moving overhead may be explained by the great ambiguity of such a signal, because this crab is immersed in an environment featuring wind-induced oscillations of the upper portion of the cord grasses which may elicit the escape response (Tomsic et al. 1993).



Neurobiology of Learning and Memory of Crustaceans

Fig. 19.3. Effect of shifting the context between training and testing sessions. (A) Animals were trained in a normal or in a striped actometer. Twenty-four hours after training, the animals were tested in the same context in which they had been trained or in the different one. (B) Left side: testing performance in normal container. Right side:  testing performance in striped container. White bars:  standard pair of untrained (U)  and trained (T) groups with the same context at training and testing sessions. Black bars: context-shift pair of untrained (U) and trained (T) groups with different context at training and testing sessions. Only the animals that were tested in the same context of the training evoked the memory. Ordinate, mean escape response SEM. *p < 0.05. **p < 0.01. For each group n = 40. From Tomsic et al. (1998), with permission from Springer.

This interpretation is supported by the fact that Pachygrapsus marmoratus, another grapsoid crab that inhabits a barren biotope compared to that of Neohelice, is by far less inclined to reduce its response to the VDS (Tomsic et al. 1993). The adaptive value of the between-species difference would be that Neohelice saves time and energy by curbing its innate tendency to escape from a passing object unlikely to be a predator, whereas Pachygrapsus keeps safe from enemies by maintaining a high response level to a passing object likely to be a predator. But under such circumstances, how does Neohelice cope with the attack of a real predator? Unlike Pachygrapsus, which inhabits rock crevices, Neohelice invests time and effort in digging its own burrow. The burrow-centered habits may have contributed to the ability of Neohelice to learn and recognize the contextual environment. Thus, the crab may recognize the familiar layout of cord grasses that surrounds its burrow. Hence, it may learn to disregard the overhead movements normally occurring in this particular context. But, if for any reason the animal travels beyond its home range, it would regain its ability to escape from similar stimuli. Thus, in Neohelice, the risk that involves the long-term waning of the escape response is counteracted by the fact that the learned inhibition is stimulus- and context specific.

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Daniel Tomsic and Héctor Maldonado The above interpretation is based on results obtained in experiments performed in the laboratory. A characteristic of these experiments is that the animal is trained inside a container from which it cannot escape. A condition where, besides learning that the stimulus is not dangerous, the animal can also learn that to escape is actually impossible, that is, an instance of learned helplessness. On the other hand, a replacement of the escape response by a freezing response following training has been described and interpreted as an instance of fear conditioning (Pereyra et al. 1999, 2000). Further field studies, like those performed by Fathala et al. (2010a, 2010b) will be necessary to be certain about the adaptive value of CSM of Neohelice (see the section Future Directions). Frequency and Number of Training Trials Determine the Type of Memory In addition to the number of training trials, memory formation is known to critically depend on the frequency of trial presentations. In different animals and using various learning tasks, short intertrial intervals (ITI), which are known as massed training, result in short or intermediate memories, whereas long intertrial intervals (spaced training) result in long-lasting memories (e.g., Tully et al. 1994, Menzel et al. 2001). This subject has been extensively investigated in Neohelice. Experiments using different frequencies and amounts of training trials revealed that massed and spaced training build up different types of memory (Fig. 19.2) (Pedreira et al. 1998). Fifteen trials of spaced training (ITI: 3 min) result in a long-lasting memory (>5 days), while a similar number of massed training trials results in a rapid reduction of the escape response that is only short lasting (few minutes). Increasing the number of trials with massed training can render an intermediate memory (1 h) induces extinction, both depending on de novo protein synthesis. Based on these findings, we proposed that the reminder duration acts as a switch guiding the memory course toward reconsolidation (short reminder) or extinction (long reminder, Pedreira and Maldonado 2003). Moreover, using cycloheximide to determine the lability state of memory at different time points, we showed that two factors, no-reinforcement during the reminder (i.e., context reexposure) and the end of context reexposure, are critical for both processes to occur. We concluded that the mismatch between what is expected and what actually occurs is the mandatory condition that leads CSM toward reconsolidation or extinction (Pedreira et al. 2004). Grounded in these and other results (e.g., Pérez-Cuesta et  al. 2007, Pérez-Cuesta and Maldonado 2009, Hepp et al. 2010), two diagnostic features emerge for the reconsolidation process. The first is the labilization of the reactivated memory, revealed by the amnesic effect of an interfering factor presented after the reminder. The second is the specificity of the reminder structure, revealed by the fact that the amnesic effect of the interference is no longer observed when the reminder’s parametrical conditions are changed. Inspired by our work with Neohelice, we designed a new paradigm to test the reconsolidation hypothesis in human declarative memory. In our experiment with verbal material (Forcato et al. 2007), the target memory was acquired by a first learning process and the interfering agent was a second learning acquired after the presentation of the reminder. In short, our experiments on reconsolidation of human declarative memory confirmed the two diagnostic features of reconsolidation found in our studies with the crab, that is, labilization of the retrieved memory, and specificity of the reminder (Forcato et al. 2009, 2010). Anatomical and Electrophysiological Identification of Neurons Supporting Context-Signal Memory The CSM of Neohelice is based on visual information. For that reason, 10 years ago we started to apply electrophysiological techniques to search for neurons in the optic neuropils that may house the memory trace. Since then, unforeseen progress has been made. Results led to the first identification in an arthropod of neurons supporting long-term visual memories, and, in addition, provided a great deal of new knowledge on the anatomical and physiological organization of the visual nervous system of decapods. Thus, before describing the neurons involved in the visual learning and memory of the crab, a brief description of the organization of the visual nervous system is needed (for general information on the visual system of crustaceans, see ­chapter 8 in this volume). Each eye of N.  granulata (Fig. 19.4) consists of about 8,000 ommatidia, which are distributed around the tip of the eyestalk except for a narrow area of cuticle located toward the medial side of the animal. Thus, the visual field of each eye subtends almost the entire panorama surrounding the animal. As is typical of decapods, each of the two optic lobes consists of three nested retinotopic neuropils plus an additional fourth retinotopic neuropil more recently discovered which, with a number of circumscribed protocerebral neuropils,



Neurobiology of Learning and Memory of Crustaceans

Fig. 19.4. Visual space maps of the optic neuropils of the crab. Please see color version of this figure in the center plate. Upper and lower panels represent frontal and lateral views (from a slightly dorsal perspective) of the right eye and its optic neuropils. Doubled vertical lines represent meridian positions across the retina at 0, 90, and 180 degrees, while thick horizontal lines represent positions in elevation at the eye equator (0 degrees), and at 40 degrees above and 20 degrees below the equator. This code of doubled vertical lines and thick horizontal lines is preserved across the three optic neuropils and the chiasmata. In the crab, the first optic chiasm inverts the order of representation corresponding to the horizontal plane (upper panel), while the second optic chiasm inverts the order corresponding to the vertical plane (lower panel). Scale is not provided because the absolute dimensions vary with the size of the animal. Abbreviations: La, lamina; Me, medulla; Lo, lobula; O Ch1, first optic chiasm; O Ch2, second optic chiasm. From Berón de Astrada et al. (2011), with permission from Wiley.

are contained in the eyestalk. These are connected to medial neuropils of the supraesophageal ganglion by discrete protocerebral tracts. From periphery to center, the three optic neuropils are traditionally called lamina ganglionaris, external medulla, and internal medulla. However, modern anatomical and developmental studies argue for them being named the lamina, medulla, and lobula, so as to conform with homologous neuropils in insects (Sinakevitch et al. 2003, Sztarker et al. 2005, Strausfeld 2009, Krieger et al. 2010). The arrangement of the optic neuropils within the eyestalk of Neohelice and its retinotopic organization are schematized in Fig. 19.4 (see color version in center plate) and shown in Fig. 19.5. The principal architectural elements are retinotopic columns intersected by layers. Each column or cartridge represents a visual sampling unit of the retina: photoreceptors from each ommatidium send their axons into a lamina optic cartridge, each of which is delineated by a rectilinear organization of tangentially directed processes. Efferent neurons extending from optic cartridges project the distal representation of visual sampling units proximally into the medulla. Deeper retinotopic organization is preserved through the medulla by its columnar neurons as they map this geometry into the lobula. In malacostracans as in insects, axons connecting the lamina with the medulla and the medulla with the lobula form chiasmata

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Daniel Tomsic and Héctor Maldonado (Fig. 19.4 and 19.5). The neural architecture, anatomical organization, and space maps of the optic lobes of Neohelice are described in detail elsewhere (Sztarker et al. 2005, 2009, Berón de Astrada et al. 2011). Below we present an overview of the lobula because of its central role in visual memory. Like the medulla, the lobula of Neohelice is a dome shaped structure, slightly elongated in the lateromedial axis (Fig. 19.4). As in the lamina and medulla, the fibroarchitectural appearance of the lobula depends on the section orientation because different cell types have their tangential processes exclusively oriented either along the anteroposterior or the lateromedial axis. Transverse sections show four strata of tangential processes oriented lateromedially. From the periphery to the center, these strata are referred to as the first to fourth lateromedial tangential layers (LMT1-LMT4) (Fig. 19.6C). These strata are separated by regions containing arborization profiles belonging to columnar elements and local interneurons, and the profiles of tangential processes running anteroposteriorly. Both LMT1 and LMT4 comprise relatively thin tangential processes, whereas LMT2 and LMT3 contain long wide-diameter tangential fibers that increase in girth toward the medial side of the neuropil (Fig. 19.6C). Intracellular staining (Fig. 19.6B) confirmed that LMT2 and LMT3 layers comprise the bistratified dendritic tree of wide-field motion-sensitive neurons termed bistratified lobula giant (B-LG) neurons, to distinguish them from the monostratified type described below. The dendritic tree of each of these neurons consists of several branches that run parallel to each other all along the lateromedial axis of the lobula. The dendrites converge toward the medial side of the neuropil into a thicker

Fig. 19.5. Overview of a 12- μ m slice of Neohelice optic neuropils stained with Bodian’s reduced-silver. Longitudinal section of the entire lobe. The visual neuropils are divided into discrete strata. The lamina (La) is connected to the medulla by the first optic chiasm (OCh1), and the medulla is connected to the lobula by the second chiasm (OCh2). The lobula provides many discrete tracts to the protocerebrum. The neuron somata are arranged as layers or clusters outside neuropils. Scale bar = 50 μ m. Modified from Sztarker et al. (2005), with permission from Wiley.



Neurobiology of Learning and Memory of Crustaceans B

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Fig. 19.6. The motion-sensitive lobula giant (LG) neurons. (A) Reconstruction based on several intracellular fills showing the anterior view of the structure of a bistratified LG neuron type 2 (B-LG2). (B)  Transverse section of the lobula showing an intracellularly stained B-LG2 with its typical bistratified branching pattern (brackets). (C) In reduced-silver impregnated lobulas, the bistratified branches can be identified as forming part of layers LMT2 and LMT3 (marked by the two central brackets) (the more distal and proximal brackets correspond to LMT1 and LMT4 respectively). The proximal branches of B-LG2 are intercepted by 14 thick processes that extend normal to this plane of section. These are the primary processes of a system of 14 monostratified LG neurons of type 1 (M-LG1), which are oriented anteroposteriorly. (D) Intracellular staining of an MLG1, revealing the thick main process running anteroposteriorly, from which secondary processes arise at right angles. The thin neurite connecting the soma to the main process (white arrow) and the descending axon from this process (black arrow) are also filled. (E) Reduced silver preparation showing the parallel arrangement of primary processes (black arrows) from several M-LG1s. From the primary processes of M-LG1s, secondary processes can be observed running parallel and close to the proximal arborizations of B-LG2. Abbreviations: Lo, lobula; LPc, lateral protocerebrum; PcT, protocerebral tract. Scale bars = 100 μ m. Modified from Sztarker et al. (2005), with permission from Wiley.

single axonal trunk that can be followed toward the midbrain (Berón de Astrada and Tomsic 2002, Sztarker et al. 2005, Medan et al. 2007). Longitudinal sections of the lobula demonstrate five strata composed of tangential processes oriented anteroposteriorly. From the periphery to the center, these are called the anteroposterior tangential levels (APT1-APT5). The strata are separated by clearer layers in which the organization of columnar elements is apparent and from which arise processes that are oriented

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Daniel Tomsic and Héctor Maldonado perpendicular to the plane of the section. Level APT4 deserves special attention as it is composed of 14 neurons, each of which possesses an exceptionally wide-diameter primary branch (8–10 μ m diameter) oriented along the anteroposterior axis (Fig. 19.6C–E). Primary branches from these 14 cells are arranged in parallel, separated from one another by approximately 35 μ m (Fig. 19.6C,E). Each primary branch provides several secondary processes that arise at right angles and thus extend lateromedially within the LMT3 stratum in close proximity to the proximal tangential processes of the B-LGNs (Figs. 19.6C). Each primary branch is connected to a prominent axon that descends through the lateral protocerebrum to reach the optic tract. The descending axons of the 14 neurons converge to form a discrete bundle. The large somata of these neurons are clustered anteromedially to the lobula, each being connected by a neurite to its primary branch (Fig. 19.6D). As will be described next, intracellular studies indicate that these cells, hereafter termed monostratified lobula giants (M-LGs), integrate the group of motion sensitive lobula giant (LG) neurons. In Vivo Physiological Characterization of Brain Interneurons Adaptive behavior in a cluttered visual environment requires that animals extract information from the visual scene. Ideally, neuronal events related to behavioral performance should be investigated while they occur in vivo, but physical accessibility and stability limit intracellular studies. We developed a preparation with Neohelice that allows the investigation of the physiological responses of individual neurons by intracellular recordings in the intact living animal (Fig. 19.1B; for methods, see Berón de Astrada et al. 2001, Medan et al. 2007). Toward this aim, the crab offers unique advantages: (1) the visual neuropils are easily accessed without damage to the animal; (2) the hard carapace provides the mechanical stability required for intracellular recordings; and (3) Neohelice has a great resistance to manipulation. Remarkably, following several hours of electrophysiological recording, the crab remains healthy and, days after the experiment, no behavioral differences are observed with respect to nontreated animals (Tomsic et al. 2003). Although crustaceans represent one of the major phyletic groups, there is little knowledge about their neural processing of visual information (see ­c hapter  8 in this volume). Thus, we started to characterize the electrophysiological properties of different types of neurons by their responses to a pulse of light or to moving stimuli delivered to the intact animal. We also identified the location and morphology of these neurons by intracellularly staining them with neurobiotin. Neurons from the lamina and medulla react more vigorously to a pulse of light than to a moving stimulus (Berón de Astrada et al. 2001, 2009, Berón de Astrada and Tomsic 2002), while the recorded neurons from the lobula are clearly tuned to respond to visual motion (Berón de Astrada and Tomsic 2002, Tomsic et al. 2003, Medan et al. 2007, Oliva et al. 2007). Because we are interested in learning and memory processes induced by the repeated presentation of a motion stimulus, the LG neurons have been the main focus of our attention.

Characterization of the Lobula Giants and Their Role in the Crab Escape Response In Neohelice, the sudden movement of a VDS above the animal elicits a strong escape reaction. We have morphologically identified and physiologically characterized four classes of LGs that responded to the same stimulus that elicits the behavioral reaction (in our early studies we called these elements movement detector neurons). Two of the classes present arborizations



Neurobiology of Learning and Memory of Crustaceans

in the lobula that are monostratified (M-LG types 1 and 2), while the other two classes are bistratified (B-LG types 1 and 2) (Fig. 19.6, Medan et al. 2007). Specific features of these neurons have been described elsewhere (Berón de Astrada and Tomsic 2002, Sztarker and Tomsic 2004, 2008, Medan et al. 2007, Oliva et al. 2007). Briefly, the response to a moving stimulus consists of a strong discharge of action potentials frequently superimposed on noisy graded potentials (Fig. 19.1B). The response to a single moving object is more intense than to the movement of the whole visual field, indicating that these neurons are tuned to object detection rather than to flow field processing. As is characteristic in all movement sensitive neurons, including those of vertebrates, the response of the LGs is relatively independent of the background intensity and the contrast between moving target and background. The response of each neuron is highly consistent on repeated stimulation, but such consistency can be obtained only when the stimuli are separated by long intervals. Repeated stimulation at intervals shorter than 10 min produces a reduction of the response (see below). The extent and location of the receptive field, as well as the directional sensitivity, varies among the different LG classes. Intracellular recordings also showed that the LGs respond to visual stimuli presented either to the ipsilateral or to the contralateral eye, thus demonstrating that processing of binocular visual information occurs at the level of the lobula (Sztarker and Tomsic 2004). In addition, three of the four classes respond to both visual and mechanical stimuli applied to areas of the body, demonstrating that the integration of multimodal information also occurs in the lobula. The general characteristics of the LGs of Neohelice coincide with those of the morphologically unidentified movement fibers largely studied by Wiersma and coworkers in different decapod species (reviewed in Wiersma et  al. 1982; see ­chapter 8 in this volume). A variety of experimental conditions that affected the level of escape, such as seasonal variations, changes in stimulus features, and whether the crab perceived stimuli monocularly or binocularly, also consistently affected the response of LG neurons in a way that closely matched the effects observed at the behavioral level (Sztarker and Tomsic 2008, 2011). The analysis showed that the firing profile of the LGs corresponds well with that of the behavioral reaction, whereas a comparison of the times taken to reach half of the maximum responses revealed that on average the neuronal reaction anticipates the behavioral response by about 120 msec (Tomsic et al. 2003, Oliva et al. 2007). The sum of evidence suggests that the LGs play a central role in the circuit that controls the escape response elicited by VDS. Lobula Giants and Their Role in Visual Learning and Memory In Neohelice, the escape response elicited by VDS declines after few stimulus presentations. As we already described, high-frequency presentation (massed training) produces rapid but short-lasting escape suppression, whereas low-frequency stimulation (spaced training) results in a much slower but enduring behavioral change. We found that the rates of reduction and recovery of the response of LGs in both massed and spaced training remarkably coincide with the changes occurring at the behavioral level. Massed training yields a fast and profound reduction of the response, which recovers completely after 15 min at both the behavioral and neuronal levels (Fig. 19.7A,C). In contrast, spaced training results in a slower and shallower reduction of the response, but the changes last longer (Fig. 19.7B,D). The change of response is specific for the moving stimulus, because the response of the neuron to a pulse of light measured before and after repeated presentation of the training stimulus is not affected (Tomsic et al. 2003). In the experiments described above, changes in the response of the LGs were assessed up to 15 min following training. Since spaced training induces a memory that lasts for several days, we investigated whether the neuronal changes observed 15 min after training persisted for at least

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Fig. 19.7. Changes in LG neurons reflect learning as well as short- and long-term memory. (A)  Fifteen trials of massed training (ITI = 0 sec) cause a rapid and deep reduction of the response of the LG which, however, recovers completely in less than 15 min (test). (B) The same amount of spaced training (ITI = 3 min) results in a slow decrease of the response which, however, endures until the test. Trials 1, 5, and 15 and test trial are shown. Each training trial comprises two cycles of stimulus movement separated by 2 sec. (C) Averaged neuronal responses as those showed in (A) (n = 10 neurons from 10 crabs) compared to averaged behavioral responses (n = 20 crabs) evoked by massed training. (D) Averaged neuronal responses as those showed in (B) (n = 12 neurons from 12 crabs) compared to averaged behavioral responses (n = 20 crabs) evoked by spaced training. Control (n = 8) refers to LGs from crabs that received only the first trial and the test trial. Although massed and spaced training induce different response changes, within each training condition the behavioral and neural performances appear identical. Responses were normalized to the first response of each recording. (E) Long-term changes in the LGs reflect long-term memory. One day after spaced training, the escape response (control n = 40 crabs; trained n = 40 crabs) and the neuronal response (control n = 40 neurons from 40 crabs; trained n = 40 neurons from 40 crabs) remain both significantly suppressed. p < 0.05 (two-sample t-test; two sides p-value). Graphs display means SEM. Modified from Tomsic et al. (2003), with permission from The Society for Neuroscience.



Neurobiology of Learning and Memory of Crustaceans

24 h after training. Thus, was long-term memory in the crab reflected by long-term change in the response of the LGs? To determine this, we trained crabs with 30 spaced trials, and 24 h later compared their behavioral and neuronal responses with those of control animals. The comparisons showed that training similarly affected the behavioral and neuronal responses (Fig. 19.7E). Several lines of evidence indicate that the changes observed in LGs do not result from changes occurring in presynaptic neurons. For example, neurons from the medulla were found not to change on repeated VDS presentations (Tomsic et al. 2003). Besides, unpublished results indicate that the long-term neuronal reduction is not accounted for by changes in the input signal, since no changes in the postsynaptic potential (PSP) response were observed. The evidence then, suggests that learning induces a change in the transfer of information from the input to the output neuronal signals (PSP to elicited spikes). The results demonstrate that long-term memory of the VDS, which is expressed as a reduction in the intensity of the escape reaction, is supported by long-term changes that take place in the giant neurons of the lobula (Tomsic et al. 2003, 2009). Recently, we found that the ability of crabs to generalize the learned stimulus into new space positions and to distinguish it from a similar but unlearned stimulus, two of the main attributes of stimulus-memory, can be completely explained by the performance of LGs (Sztarker and Tomsic 2011). As already mentioned, CSM can only be evoked in the training context, implying an association between a CM and an SM. Therefore, animals seem to store two visual components of the learned experience, one related to the stimulus (SM) and one related to the context (CM). In the previously described experiments, the response of LGs from trained crabs was assessed while keeping the contextual environment constant between training and testing. Hence, an important question that remained was whether the effect of changing the context in the testing session would be reflected by the performance of the LGs. In other words, do these neurons store the entire CSM trace or only the SM component? To test this, we evaluated the response of LGs to the VDS in animals located in the same or in a different visual context compared with where the memory had been acquired. We found that LGs do not support the visual context-memory component. Our results provided the first physiological evidence that memory traces regarding “what” and “where” are stored separately in the arthropod brain (Sztarker and Tomsic 2011).

FUTURE DIRECTIONS Understanding the way in which a brain, even a small one, is organized and functions to produce behavior is still a huge challenge. The following are major pending questions in the study of learning and memory for which crustaceans offer good opportunities of research. Memory, expressed as relatively endurable changes of behavior as a result of experience, implicates sustained modifications of neural connectivity. We are just beginning to understand the mechanisms underlying it. Most of what we know about the intrinsic and synaptic changes that neurons are capable of derives from studies performed in highly reduced preparations, such as brain slices or cell culture. However, neurons naturally operate within a brain, which in turn is deeply inserted in and connected with a living acting animal and its environment. How close are the mechanisms of neuronal plasticity that we discover in the highly artificial environment of reduced preparations to those occurring in the intact animal when learning? The strong carapace of crabs, which easily allows firmly holding them, together with the accessibility to important portions of their brain and their ability to withstand experimental manipulation, provide excellent opportunities to investigate changes that occur in single identified neurons

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Daniel Tomsic and Héctor Maldonado in a completely intact animal during learning and memory (Tomsic et al. 2003, Sztarker and Tomsic 2011). Studies of the neuronal mechanisms of learning and memory are almost exclusively performed in the laboratory, under highly controlled conditions, where animals are usually trained to learn single stimuli relationships in rather artificial setups. While there are clear methodological advantages to this approach, it may also lead to a misapprehension of the real abilities that animals are capable of when performing in their natural environment. In other words, how much of the restricted cognitive abilities ascribed to animals result from our dominant perspective of studying the neurobiology of learning and memory in the laboratory? The LG neurons, which proved to support complex stimulus-memory attributes (Sztarker and Tomsic 2011), and which are thought to play a central role in controlling visually guided behaviors (e.g., Oliva et al. 2007, Sztarker and Tomsic 2008), project their axon along the protocerebral tract (Medan et  al. 2007). Thus, it is technically feasible to record the activity of the LGs in the behaving animal using chronically implanted electrodes (Wiersma et al. 1982). Moreover, given the size and strength of crabs, electrophysiological recording could be made in the field using miniature data loggers. This, in combination with the possibility offered by crabs to thoroughly investigate their behavior using video recording in the natural environment (see ­chapter 18 in this volume), provide an opportunity to address the type of questions raised above. The possibilities emerging from combining laboratory and field studies in crabs to understand the neural control of behavior has been discussed in a recent review (Hemmi and Tomsic 2012). Interestingly, studies in crayfish (Hazlett et al. 2002) and crabs (Roudez et al. 2008) demonstrated significantly greater learning abilities for invasive species when compared with native relatives. It has been argued that higher learning capacities may account in part for the success of invasive species (Weis 2010, Wright et al. 2010). Therefore, knowing the abilities of different crustacean species for learning would provide more insight into the causes and consequences of species invasions.

SUMMARY AND CONCLUSIONS Learning and memory abilities are not limited to big-brained animals, but rather these are essential adaptive features of all animals that actively explore their environment. Studies on the neurobiology of learning and memory using the crab Neohelice as an experimental model highlight the value of using decapods for investigating general principles of the neural control of behavior. For instance, studies on memory reconsolidation in the crab led to new interpretations on the way in which memories are stored and upgraded (Pedreira et  al. 2002, Pedreira and Maldonado 2003). Remarkably, these experiments inspired experiments in human memory, which confirmed the basic principles derived from the studies made in the crab (Forcato et al. 2007, 2009, 2010). Another example of the advantages offered by some decapods for neurophysiological studies is the possibility of recording the activity of central neurons in intact animals. In fact, recording brain activity in vivo during learning is fundamental to understanding how memories are formed. However, intracellular recordings are rarely achieved in intact, behaving organisms. It is notable that in Neohelice, neuronal changes can be assessed by intracellular recordings in a living animal at the same time it is learning. Moreover, after the recording, the crab remains healthy, and several days after the experiment, no behavioral differences are observed with respect to nontreated animals. Thus, the visual memory abilities of crabs, their relatively simple and accessible nervous system, and the recording stability that can be achieved with their



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neurons provide an opportunity for uncovering neurophysiological and molecular events that occur in identifiable neurons during learning.

ACKNOWLEDGMENTS We thank Sergio Nemirovsky, Julieta Sztarker, and Fiorella Magani for fruitful comments and correction of the manuscript.

REFERENCES Abramson, C.I., and R.D. Feinman. 1987. Operant punishment of eye elevation in the green crab, Carcinus maenas. Behavioral and Neural Biology 48:259–277. Abramson, C.I., and R.D. Feinman. 1988. Classical conditioning of the eye withdrawal reflex in the green crab. Journal of Neuroscience 8:2907–2912. Abramson, C.I., and R.D. Feinman. 1990. Lever-press conditioning in the crab. Physiology and Behavior 48:267–272. Abramson, C.I., P.M. Armstrong, R.A. Feinman, and R.D. Feinman. 1988. Signaled avoidance in the eye withdrawal reflex of the green crab. Journal of Experimental Analysis of Behavior 50:483–492. Aggio, J., A. Rakitin, and H. Maldonado. 1996. Serotonin-induced short- and long-term sensitization in the crab Chasmagnathus. Pharmacology Biochemistry and Behavior 53:441–448. Appleton, T., and J.L. Wilkens. 1990. Habituation, sensitization and the effect of serotonin on the eyestalk withdrawal reflex of Cancer magister. Comparative Biochemistry and Physiology 97A:159–163. Basil, J., and D. Sandeman. 2000. Crayfish (Cherax destructor) use tactile cues to detect and learn topographical changes in their environment. Ethology 106:247–259. Berón de Astrada, M., and H. Maldonado. 1999. Two related forms of long-term habituation in the crab Chasmagnathus are differentially affected by scopolamine. Pharmacology Biochemistry and Behavior 63:109–118. Berón de Astrada, M., J. Sztarker, and D. Tomsic. 2001. Visual interneurons of the crab Chasmagnathus studied by intracellular recordings in vivo. Journal of Comparative Physiology A 187:37–44. Berón de Astrada, M., and D. Tomsic. 2002. Physiology and morphology of visual movement detector neurons in a crab (Decapoda: Brachyura). Journal of Comparative Physiology A 188:539–551. Berón de Astrada, M., J.C. Tuthill, and D. Tomsic. 2009. Physiology and morphology of sustaining and dimming neurons of the crab Chasmagnathus granulatus (Brachyura: Grapsidae). Journal of Comparative Physiology A 195:791–798. Berón de Astrada, M., V. Medan, and D. Tomsic. 2011. How visual space maps in the optic neuropils of a crab. Journal of Comparative Neurology 519:1631–1639. Bruner, J., and D. Kennedy. 1970. Habituation: occurrence at a neuromuscular junction. Science 169:92–94. Brunner, D., and H. Maldonado. 1988. Habituation in the crab Chasmagnathus granulatus: effect of morphine and naloxone. Journal of Comparative Physiology A 162:687–694. Cannicci, S., R.K. Ruwa, and M. Vannini. 1997. Homing experiment in the tree-climbing crab Sesarma leptosoma (Decapoda, Grapsidae). Ethology 103:935–944. Cannicci, S., S. Fratini, and M. Vannini. 1999. Short range homing in fiddler crabs (Ocypodidae, genus Uca): a homing mechanism not based on local visual landmarks. Ethology 105:867–880. Cannicci, S., C. Barelli, and M. Vannini. 2000. Homing in the swimming crab Thalamita crenata: a mechanism based on underwater landmark memory. Animal Behaviour 60:203–210. Cronin, T.W. 2006. Stomatopods. Current Biology 16:235–236. Cuadras, J., E. Vila, and J. Balasch. 1978. T-maze shock avoidance in the hermit crab Dardanus arrosor. Revista Española de Fisiología 34:273–276.

529

530

Daniel Tomsic and Héctor Maldonado Chittka, L., and J. Niven. 2009. Are bigger brains better? Current Biology 19:995–1008. Delorenzi, A., M.E. Pedreira, A. Romano, S.I. Garcia, C.J. Pirola, V.E. Nahmod, and H. Maldonado. 1996. Angiotensin II enhances long-term memory in the crab Chasmagnathus. Brain Research Bulletin 41:211–220. Delorenzi, A., B. Dimant, L. Frenkel, V.E. Nahmod, D.R. Nässel, and H. Maldonado. 2000. High environmental salinity induces memory enhancement and increases levels of brain angiotensin-like peptides in the crab Chasmagnathus granulatus. Journal of Experimental Biology 203:3369–3379. Denti, A., B. Dimant, and H. Maldonado. 1988. Passive avoidance learning in the crab Chasmagnathus granulatus. Physiology and Behavior 43:317–320. Dimant, B., and H. Maldonado. 1992. Habituation and associative learning during exploratory behavior of the crab Chasmagnathus. Journal of Comparative Physiology A 170:749–759. Dimant, B., A. Rossen, and G. Hermitte. 2002. CS-US delay does not impair appetitive conditioning in Chasmagnathus. Behavioral Processes 60:1–14. Dudai, Y. 2008. Memory from A to Z: keywords, concepts, and beyond. Oxford University Press, New York. Eisenstein, E.M., and P.J. Mill. 1965. Role of the optic ganglia in learning in the crayfish Procambarus clarkii (Girard). Animal Behaviour 13:561–565. Fathala, M. del V., L. Iribarren, M.C. Kunert, and H. Maldonado. 2010a. A field model of learning: 1. Short-term memory in the crab Chasmagnathus granulatus. Journal of Comparative Physiology A 196:61–75. Fathala, M. del V., M.C. Kunert, and H. Maldonado. 2010b. A field model of learning: 2. Long-term memory in the crab Chasmagnathus granulatus. Journal of Comparative Physiology A 196:77–84. Federman, N., M.S. Fustiñana, and A. Romano. 2009. Histone acetylation is recruited in consolidation as a molecular feature of stronger memories. Learning and Memory 16:600–606. Feld, M., B. Dimant, A. Delorenzi, O. Coso, and A. Romano. 2005. Phosphorylation of extra-nuclear ERK/MAPK is required for long-term memory consolidation in the crab Chasmagnathus. Behavioral Brain Research 158:251–261. Fernandez-Duque, E., C. Valeggia, and H. Maldonado. 1992. Multitrial inhibitory avoidance learning in the crab Chasmagnathus. Behavioral and Neural Biology 57:189–197. Forcato, C., V.L. Burgos, P.F. Argibay, V.A. Molina, M.E. Pedreira, and H. Maldonado. 2007. Reconsolidation of declarative memory in humans. Learning and Memory 14:295–303. Forcato, C., P.F. Argibay, M.E. Pedreira, and H. Maldonado. 2009. Human reconsolidation does not always occur when a memory is retrieved: the relevance of the reminder structure. Neurobiology of Learning and Memory 91:50–57. Forcato, C., M.L. Rodríguez, M.E. Pedreira, and H. Maldonado. 2010. Reconsolidation in humans opens up declarative memory to the entrance of new information. Neurobiology of Learning and Memory 93:77–84. Frenkel, L., R. Freudenthal, A. Romano, V.E. Nahmod, H. Maldonado, and A. Delorenzi. 2002. Angiotensin II and the transcription factor Rel/NF-κB link environmental water shortage with memory improvement. Neuroscience 115:1079–1087. Freudenthal, R., F. Locatelli, G. Hermitte, H. Maldonado, C. Lafourcade, A. Delorenzi, and A. Romano. 1998. κ-B like DNA-binding activity is enhanced after spaced training that induces long-term memory in the crab Chasmagnathus. Neuroscience Letters 242:143–146. Freudenthal, R., and A. Romano. 2000. Participation of Rel/NF-κB transcription factors in long-term memory in the crab Chasmagnathus. Brain Research 855:274–281. Fricke, R.A. 1984. Development of habituation in the crayfish due to selective weakening of electrical synapses. Brain Research 322:139–143. Fustiñana, M.S., P. Ariel, N. Federman, R. Freudenthal, and A. Romano. 2010. Characterization of the beta amyloid precursor protein-like gene in the central nervous system of the crab Chasmagnathus. Expression during memory consolidation. BMC Neuroscience 11:109. Gherardi, F., E. Tricarico, and J. Atema. 2005. Unraveling the nature of individual recognition by odor in hermit crabs. Journal of Chemical Ecology 31:2877–2896.



Neurobiology of Learning and Memory of Crustaceans

Giurfa, M., S. Zhang, A. Jenett, R. Menzel, and M.V Srinivasan. 2001. The concepts of ‘sameness’ and ‘difference’ in an insect. Nature 410:930–933. Glantz, R.M. 1974. The visually evoked defense reflex of the crayfish: habituation, facilitation, and the influence of picrotoxin. Journal of Neurobiology 5:263–280. Godoy, A.M., and H. Maldonado. 1995. Modulation of the escape response by [D-Ala2]Met-enkephalin in the crab Chasmagnathus. Pharmacology Biochemistry and Behavior 50:445–451. Hazlett, B.A. 1969. “Individual” recognition and agonistic behaviour in Pagurus bernhardus. Nature 222:268–269. Hazlett, B.A., A. Burba, F. Gherardi, and P. Acquistapace. 2002. Invasive species of crayfish use a broader range of predation-risk cues than native species. Biological Invasions 5:223–228. Hemmi, J.M., and T. Merkle. 2009. High stimulus specificity characterizes anti-predator habituation under natural conditions. Proceedings of the Royal Society London B: Biological Sciences 276:4381–4388. Hemmi, J.M., and D. Tomsic. 2012. The neuroethology of escape in crabs: from sensory ecology to neurons and back. Current Opinion in Neurobiology 22:194–200. Hepp, Y., L.M. Pérez-Cuesta, H. Maldonado, and M.E. Pedreira. 2010. Extinction memory in the crab Chasmagnathus: recovery protocols and effects of multi-trial extinction training. Animal Cognition 13:391–403. Hermitte, G., M.E. Pedreira, D. Tomsic, and H. Maldonado. 1999. Context shift and protein synthesis inhibition disrupt long-term habituation after spaced, but not massed, training in the crab Chasmagnathus. Neurobiology of Learning and Memory 71:34–49. Hiatt, R.W. 1948. The biology of the lined shore crab, Pachygrapsus crassipes Randall. Pacific Science 2:135–213. Hirsh, R., and C.A.G. Wiersma. 1977. The effect of the spacing of background elements upon optomotor memory responses in the crab: the influence of adding or deleting features during darkness. Journal of Experimental Biology 66:33–46. Hoyle, G. 1976. Learning of leg position by the ghost crab Ocypode ceratophthalma. Behavioral Biology 18:147–163. Hughes, R.N., and N. O’Brien. 2001. Shore crabs are able to transfer learned handling skills to novel prey. Animal Behaviour 61:711–714. Hwang, J.S., J.H. Costello, and J.R. Stickler. 1994. Copepod grazing in turbulent flow: elevated foraging behavior and habituation of escape responses. Journal of Plankton Research 16:421–431. Jackson, N.W., and R.W. Elwood. 1989. Memory of information gained during shell investigation by the hermit crab, Pagurus bernhardus. Animal Behaviour 37: 529–534. Kawai, N., R. Kono, and S. Sugimoto. 2004. Avoidance learning in the crayfish (Procambarus clarkii) depends on the predatory imminence of the unconditioned stimulus: a behavior systems approach to learning in invertebrates. Behavioral Brain Research 150:229–237. Krasne, F.B., and J.S. Bryan. 1973. Habituation: regulation through presynaptic inhibition. Science 182:590–592. Krieger, J., R.E. Sandeman, D.C. Sandeman, B.S. Hansson, and S. Harzsch. 2010. Brain architecture of the largest living land arthropod, the giant robber crab Birgus latro (Crustacea, Anomura, Coenobitidae): evidence for a prominent central olfactory pathway? Frontiers in Zoology 7:25. Lagerspetz, K.Y., and L Kivivuori. 1970. The rate and retention of the habituation of the shadow reflex in Balanus improvisus (Cirripedia). Animal Behaviour 18:616–620. Locatelli, F., C. LaFourcade, H. Maldonado, and A. Romano. 2001. Characterisation of cAMP-dependent protein kinase isoforms in the brain of the crab Chasmagnathus. Journal of Comparative Physiology B 171:33–40. Locatelli, F., H. Maldonado, and A. Romano. 2002. Two critical periods for cAMP-dependent protein kinase activity during long-term memory consolidation in the crab Chasmagnathus. Neurobiology of Learning and Memory 77:234–249. Locatelli, F., and A. Romano. 2005. Differential activity profile of cAMP-dependent protein kinase isoforms during long-term memory consolidation in the crab Chasmagnathus. Neurobiology of Learning and Memory 83:232–242.

531

532

Daniel Tomsic and Héctor Maldonado Lozada, M., A. Romano, and H. Maldonado. 1988. Effect of morphine and naloxone on a defensive response of the crab Chasmagnathus granulatus. Pharmacology Biochemistry and Behavior 30:635–640. Lozada, M., A. Romano, and H. Maldonado. 1990. Long-term habituation to a danger stimulus in the crab Chasmagnathus granulatus. Physiology and Behavior 47:35–41. Luppi, T., C. Bas, A. Méndez Casariego, M. Albano, J. Lancia, M. Kittlein, A. Rosenthal, N. Farías, E. Spivak, and O. Iribarne. 2013. The influence of habitat, season and tidal regime in the activity of the intertidal crab Neohelice (=Chasmagnathus) granulata. Helgoland Marine Research 67:1–15. Maldonado, H. 2002. Crustaceans as models to investigate memory illustrated by extensive behavioral and physiological studies in Chasmagnathus. Pages 314–327 in K. Wiese, editor. The crustacean nervous system. Springer-Verlag, Berlin, Heidelberg. Maldonado, H., A. Romano, and M. Lozada. 1989. Opioid action on response level to a danger stimulus in the crab (Chasmagnathus granulatus). Behavioral Neuroscience 103:1139–1143. Maldonado, H., A. Romano, and D. Tomsic. 1997. Long-term habituation (LTH) in the crab Chasmagnathus: a model for behavioral and mechanistic studies of memory. Brazilian Journal of Medical and Biological Research 30:813–826. Marchand, A.R., and W.J.P. Barnes. 1992. Correlates of habituation of a polysynaptic reflex in crayfish in vivo and in vitro. European Journal of Neuroscience 4:521–532. Medan, V., D. Oliva, and D. Tomsic. 2007. Characterization of lobula giant neurons responsive to visual stimuli that elicit escape behaviors in the crab Chasmagnathus. Journal of Neurophysiology 98:2414–2428. Menzel, R., G. Manz, and U. Greggers. 2001. Massed and spaced learning in honeybees: the role of CS, US, the intertrial interval, and the test interval. Learning and Memory 8:198–208. Merlo, E., and A. Romano. 2008. Memory extinction entails the inhibition of the transcription factor NF-κB. PLoS ONE 3:e3687. Merlo, E., R. Freudenthal, and A. Romano. 2002. The IκB kinase inhibitor sulfasalazine impairs long-term memory in the crab Chasmagnathus. Neuroscience 112:161–172. Merlo, E., R. Freudenthal, H. Maldonado, and A. Romano. 2005. Activation of the transcription factor NF-κB by retrieval is required for long-term memory reconsolidation. Learning and Memory 12:23–29. Micheli, F. 1995. Behavioural plasticity in prey-size selectivity of the blue crab Callinectes sapidus feeding on bivalve prey. Journal of Animal Ecology 64:63–74. Morris, R.G., J. Inglis, J.A. Ainge, H.J. Olverman, J. Tulloch, Y. Dudai, and P.A. Kelly. 2006. Memory reconsolidation: sensitivity of spatial memory to inhibition of protein synthesis in dorsal hippocampus during encoding and retrieval. Neuron 50:479–489. Nader, K., G.E. Schafe, and J.E. Le Doux. 2000. Fear memories require protein synthesis in the amygdala for reconsolidation after retrieval. Nature 406:722–726. Oliva, D., V. Medan, and D. Tomsic. 2007. Escape behavior and neuronal responses to looming stimuli in the crab Chasmagnathus granulatus (Decapoda: Grapsidae). Journal of Experimental Biology 210:865–880. Pedreira, M.E., and H. Maldonado. 2003. Protein synthesis subserves reconsolidation or extinction depending on reminder duration. Neuron 38:863–869. Pedreira, M.E., B. Dimant, D. Tomsic, L.A. Quesada-Allue, and H. Maldonado. 1995. Cycloheximide inhibits context memory and long-term habituation in the crab Chasmagnathus. Pharmacology Biochemistry and Behavior 52:385–395. Pedreira, M.E., B. Dimant, and H. Maldonado. 1996. Inhibitors of protein and RNA synthesis block context memory and long-term habituation in the crab Chasmagnathus. Pharmacology Biochemistry and Behavior 54:611–617. Pedreira, M.E., A. Romano, D. Tomsic, M. Lozada, and H. Maldonado. 1998. Massed and spaced training build up different components of long-term habituation in the crab Chasmagnathus. Animal Learning and Behavior 26:34–45. Pedreira, M.E., L.M. Pérez-Cuesta, and H. Maldonado. 2002. Reactivation and reconsolidation of long-term memory in the crab Chasmagnathus: protein synthesis requirement and mediation by NMDA-type glutamatergic receptors. Journal of Neuroscience 22:8305–8311. Pedreira, M.E., L.M. Pérez-Cuesta, and H. Maldonado. 2004. Mismatch between what is expected and what actually occurs triggers memory reconsolidation or extinction. Learning and Memory 11:579–585.



Neurobiology of Learning and Memory of Crustaceans

Pereyra, P., H.O. De La Iglesia, and H. Maldonado. 1996. Training-to-testing intervals different from 24 h impair habituation in the crab Chasmagnathus. Physiology and Behavior 59:19–25. Pereyra, P., M. Saraco, and H. Maldonado. 1999. Decreased response or alternative defensive strategy in escape: two novel types of long-term memory in the crab Chasmagnathus. Journal of Comparative Physiology A 184:301–310. Pereyra, P., E. González Portino, and H. Maldonado. 2000. Long-lasting and context-specific freezing preference is acquired after spaced repeated presentations of a danger stimulus in the crab Chasmagnathus. Neurobiology of Learning and Memory 74:119–134. Pérez-Cuesta, L.M., Y. Hepp, M.E. Pedreira, and H. Maldonado. 2007. Memory is not extinguished along with CS presentation but within a few seconds after CS-offset. Learning and Memory 14:101–108. Pérez-Cuesta, L.M., and H. Maldonado. 2009. Memory reconsolidation and extinction in the crab: mutual exclusion or coexistence? Learning and Memory 16:714–721. Rakitin, A., D. Tomsic, and H. Maldonado. 1991. Habituation and sensitization to an electrical shock in the crab Chasmagnathus: effect of background illumination. Physiology and Behavior 50:477–487. Rankin, C.H. 2009. Introduction to special issue of neurobiology of learning and memory on habituation. Neurobiology of Learning Memory 92:125–126. Romano, A., M. Lozada, and H. Maldonado. 1990. Effect of naloxone pretreatment on habituation in the crab Chasmagnathus granulatus. Behavioral and Neural Biology 53:113–122. Romano, A., M. Lozada, and H. Maldonado. 1991. Nonhabituation processes affect stimulus specificity of response habituation in the crab Chasmagnathus granulatus. Behavioral Neuroscience 105:542–552. Romano, A., A. Delorenzi, M.E. Pedreira, D. Tomsic, and H. Maldonado. 1996a. Acute administration of a permeant analog of cAMP and a phosphodiesterase inhibitor improve long-term habituation in the crab Chasmagnathus. Behavioral Brain Research 75:119–125. Romano, A., F. Locatelli, A. Delorenzi, M.E. Pedreira, and H. Maldonado. 1996b. Effects of activation and inhibition of cAMP-dependent protein kinase on long-term habituation in the crab Chasmagnathus. Brain Research 735:131–140. Romano, A., F. Locatelli, R. Freudenthal, E. Merlo, M. Feld, P. Ariel, D. Lemos, N. Federman, and M.S. Fustiñana. 2006. Lessons from a crab: molecular mechanisms in different memory phases of Chasmagnathus. Biological Bulletin 210:280–288. Roudez, R.J., T. Glover, and J.S. Weis. 2008. Learning in an invasive and a native predatory crab. Biological Invasions 10:1191–1196. Sakai, K., M. Turkay, and S.L. Yang. 2006. Revision of the Helice/Chasmagnathus complex (Crustacea: Decapoda: Brachyura). Abhandlungen der Senckenbergischen Naturforschenden Gesellschaft 565:1–76. Schöne, H. 1961. Complex behavior. Pages 455–520 in T.H. Waterman, editor. The physiology of Crustacea, Vol. 2, Sense organs, integration, and behavior. Academic Press, New York. Shirinyan, D., T. Teshiba, K. Taylor, P. O’Neill, S.C. Lee, and F.B. Krasne. 2006. Rostral ganglia are required for induction but not expression of crayfish escape reflex habituation: role of higher centers in reprogramming low-level circuits. Journal of Neurophysiology 95:2721–2724. Sinakevitch, I., J.K. Douglass, G. Scholtz, R. Loesel, and N.J. Strausfeld. 2003. Conserved and convergent organization in the optic lobes of insects and isopods, with reference to other crustacean taxa. Journal of Comparative Neurology 467:150–172. Spivak, E. 2010. The crab Neohelice (= Chasmagnathus) granulata: an emergent animal model from emergent countries. Helgoland Marine Research 3:149–154. Strausfeld, N.J. 2009. Brain organization and the origin of insects: an assessment. Proceedings of the Royal Society London B: Biological Sciences 276:1929–1937. Sztarker, J., and D. Tomsic. 2004. Binocular visual integration in the crustacean nervous system. Journal of Comparative Physiology A 190:951–962. Sztarker, J., and D. Tomsic. 2008. Neuronal correlates of the visually elicited escape response of the crab Chasmagnathus upon seasonal variations, stimuli changes and perceptual alterations. Journal of Comparative Physiology A 194:587–596.

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Daniel Tomsic and Héctor Maldonado Sztarker, J., and D. Tomsic. 2011. Brain modularity in arthropods: Individual neurons that support “what” but not “where” memories. Journal of Neuroscience 31:8175–8180. Sztarker, J., N.J. Strausfeld, and D. Tomsic. 2005. Organization of optic lobes that support motion detection in a semiterrestrial crab. Journal of Comparative Neurology 493:396–411. Sztarker, J., N.J. Strausfeld, D. Andrew, and D. Tomsic. 2009. Neural organization of first optic neuropils in the littoral crab Hemigrapsus oregonensis and the semiterrestrial species Chasmagnathus granulatus. Journal of Comparative Neurology 513:129–150. Tomsic, D., and H. Maldonado. 1990. Central effect of morphine pretreatment on short- and long-term habituation to a danger stimulus in the crab Chasmagnathus. Pharmacology Biochemistry and Behavior 36:787–793. Tomsic, D., H. Maldonado, and A. Rakitin. 1991. Morphine and GABA: effects on perception, escape response and long-term habituation to a danger stimulus in the crab Chasmagnathus. Brain Research Bulletin 26:699–706. Tomsic, D., V. Massoni, and H. Maldonado. 1993. Habituation to a danger stimulus in two semiterrestrial crabs. Ontogenic, ecological and opioid correlates. Journal of Comparative Physiology A 173:621–633. Tomsic, D., B. Dimant, and H. Maldonado. 1996. Age related deficits of long-term memory in Chasmagnathus. Journal of Comparative Physiology A 178:139–146. Tomsic, D., M.E. Pedreira, G. Hermitte, A. Romano, and H. Maldonado. 1998. Context-US association as a determinant of long-term habituation in the crab Chasmagnathus. Animal Learning and Behavior 26:196–209. Tomsic, D., M. Berón de Astrada, and J. Sztarker. 2003. Identification of individual neurons reflecting short- and long-term visual memory in an arthropod. Journal of Neuroscience 23:8539–8546. Tomsic, D., M. Berón de Astrada, J. Sztarker, and H. Maldonado. 2009. Behavioral and neuronal attributes of short- and long-term habituation in the crab Chasmagnathus. Neurobiology of Learning and Memory 92:176–182. Troncoso, J., and H. Maldonado. 2002. Two related forms of memory in the crab Chasmagnathus are differentially affected by NMDA receptor antagonists. Pharmacology Biochemistry and Behavior 72:251–265. Tully, T., T. Preat, S.C. Boynton, and M. Del Vecchio. 1994. Genetic dissection of consolidated memory in Drosophila. Cell 79:35–47. Valeggia, C., E. Fernandez-Duque, and H. Maldonado. 1989. Danger stimulus-induced analgesia in the crab Chasmagnathus granulatus. Brain Research 481:304–308. Vannini, M., and S. Cannicci. 1995. Homing behaviour and possible cognitive maps in crustacean decapods. Journal of Experimental Marine Biology and Ecology 193:67–91. Weis, J.S., 2010. The role of behavior in the success of invasive crustaceans. Marine and Freshwater Behaviour and Physiology 43:83–98. Wiersma, C.A.G., J.L.M. Roach, and R.M. Glantz. 1982. Neural integration in the optic system. Pages 1–31 in D.C. Sandeman, and H.L. Atwood, editors. The biology of the Crustacea, Vol. 4, Neural integration and behavior. Academic Press, New York. Wright, T.F., J.R. Eberhard, E.A. Hobson, M.L. Avery and M.A. Russello. 2010. Behavioral flexibility and species invasions: the adaptive flexibility hypothesis. Ethology Ecology and Evolution 22:393–404. Zeil, J., and J.M. Hemmi. 2006. The visual ecology of fiddler crabs. Journal of Comparative Physiology A 192:1–25.

20 CRUSTACEANS AS MODEL SYSTEMS FOR TEACHING NEUROSCIENCE: PAST, PRESENT, AND FUTURE

Bruce R. Johnson, Robert A. Wyttenbach, and Ronald R. Hoy

Abstract In addition to their role as important preparations for research, crustaceans have served as model systems for teaching basic principles of signal transmission in nervous systems, particularly in student laboratory classes. We survey published and other easily available crustacean preparations used in neuroscience teaching, focusing on neuromuscular properties, synaptic transmission, neuronal excitability, sensory physiology, heart, respiratory, and intestinal muscle control, and neurogenesis. We also briefly describe select computer simulations, behavioral exercises, and commercially available exercises that contribute to neuroscience teaching, and insect neurobiology exercises that are often complementary. Finally we suggest future directions for development of crustacean teaching exercises.

INTRODUCTION Crustaceans are excellent models with which to teach physiology of nervous systems and introduce students to electrophysiological techniques. These preparations come from a long history of research, mainly concentrated on decapods (crabs, lobsters, crayfish), that continues to the present (Florey 1990, Wiese 2002a, 2002b), and includes this volume. There are advantages of using invertebrates instead of vertebrates in teaching laboratories (Deyrup-Olsen and Linder 1991): (1) They often illustrate a principle more clearly than vertebrate preparations because of the relatively small number of neurons in the nervous system, many of which are identifiable

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Bruce R. Johnson, Robert A. Wyttenbach, and Ronald R. Hoy across preparations. (2) They can survive and remain physiologically active at room temperature for long periods without supplemental oxygen. (3) They are generally less expensive than vertebrates. (4) They are not subject to the same governmental regulations regarding purchasing, storing, and dissecting as are vertebrates. (5) Finally, crayfish in particular are farmed commercially for fishing and food, so their removal from the environment does not damage local ecosystems. Most crustacean teaching exercises came from neurobiologists who used the preparations in their research laboratories and brought them into teaching laboratories (Parfitt 2002, Atwood 2008). For example, some versions of exercises on crustacean neuromuscular junctions (NMJs) stem from Harold Atwood’s physiology class at the University of Toronto beginning in 1965 (Atwood and Parnas 1968, Baierlein et  al. 2011). One of us (RRH) studied development and regeneration of the crayfish NMJ for his dissertation work and later developed this preparation for the Crawdad teaching project (Wyttenbach et al. 1999). This chapter surveys published and other easily available examples of crustacean preparations used in teaching neuroscience. We focus on teaching exercises rather than on the research that led to their development; the original research can be found in other chapters in this and related volumes (Wiese 2002a, 2002b). This overview is intended to inspire further development and modernization of old and possibly forgotten teaching preparations and to suggest material for future educational development.

EARLY DEVELOPMENTS Two early laboratory manuals were influential in development of crustacean teaching exercises: the Laboratory Exercises in Invertebrate Physiology manuals (Welsh and Smith 1949, 1960, Welsh et  al. 1968)  and the Experiments in Physiology and Biochemistry compilation (edited by G.A. Kerkut, starting in 1968). We start with an overview of these early efforts; specific exercises are described in more detail in the next section. The first edition of Laboratory Exercises in Invertebrate Physiology (Welsh and Smith 1949) has a remarkably comprehensive offering of crustacean neurobiology exercises developed for a semester-long physiology course at the Harvard Biological Laboratories. This edition included exercises on the pharmacological and neural control of decapod heart rates, facilitation of tension and contraction rate in crayfish leg muscle and claw opener muscle, interaction of excitatory and inhibitory innervation on contraction strength in decapod muscle, and extracellular recordings of crayfish caudal photoreceptor and responses to stimulation of tactile sensilla. The second edition (Welsh and Smith 1960) updated these exercises and introduced important new ones, including stimulus-response properties of the crayfish abdominal muscle receptor organs (MROs), measurement of the conduction velocity of action potentials (APs) from axons in decapod leg nerves and giant axons in the ventral nerve cord, and the effects of toxins and transmitters on spontaneous activity in the ventral nerve cord. This edition also introduced intracellular recording of muscle resting and synaptic potentials. In the first two editions, descriptions are short, with limited detail; much is purposely left unexplained so that students take responsibility for self-motivated exploration (Welsh and Smith 1949). The third edition (Welsh et al. 1968) updated earlier exercises and described protocols more fully. This edition suggested the crayfish abdominal superficial flexor muscle as a good preparation in which to match spontaneous APs of motor nerve with synaptic potentials of muscles but did not develop the preparation further. A new exercise characterizing location and response properties of sensory interneurons in crayfish circumesophageal connectives and ventral nerve cord was added. This edition also added an appendix on electrophysiological equipment and techniques that is still helpful today. The first and fifth volumes of the Experiments in Physiology and Biochemistry series contain exercises similar to those found in Welsh et al. (1968), but described in more detail. These include the innervation, synaptic physiology, and pharmacology of the neuromuscular junction of the crayfish abdominal extensor muscle (Atwood and Parnas 1968); inhibitory-excitatory



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motor neuron interactions controlling claw muscle contraction (Hoyle 1968); changes in sign and amplitude of postsynaptic potentials under different nerve stimulation protocols (Hoyle 1968); neural control and pharmacology of the crayfish heart (Florey 1968a, 1968b); and a detailed examination of the crayfish caudal photoreceptor (Hermann 1972). There are also new exercises on the pharmacology of crayfish hindgut activity (Florey 1968c) and the excitability of lobster giant axons (Hoekman and Dettbarn 1972).

THE CRUSTACEAN TOOLBOX FOR TEACHING NEUROBIOLOGY Thus, we find that most of the crustacean neurophysiology exercises commonly used today were present in the teaching literature nearly 40 years ago. In this section, we briefly describe published and easily available laboratory exercises that use crustaceans to examine nervous system structure and function. While our focus is on exercises that address physiology, many of these also incorporate simple staining to examine anatomical organization of nerve innervation patterns. See Fig. 20.1 for an annotated summary of selected crustacean physiology exercises that we describe below.

Fig. 20.1. Survey of selected student exercises highlighting crustaceans in laboratory teaching.

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Bruce R. Johnson, Robert A. Wyttenbach, and Ronald R. Hoy Neuromuscular Physiology and Synaptic Transmission Most published crustacean lab exercises address neural control of muscle activity in decapods in a variety of neuromuscular preparations. The major learning goals address the transformation of the neural code into muscle contraction, patterns of muscle innervation, excitatory and inhibitory control of contraction, pharmacology of the nerve-muscle synapse, and basic synaptic properties including synaptic integration and short-term synaptic dynamics. Leg and Cheliped The four neuromuscular physiology exercises in Welsh and Smith (1949) show how students can use simple techniques to examine important aspects of neuromuscular transformation and synaptic dynamics. Students use methylene blue staining to see that small numbers of axons innervate leg muscles and that muscle fibers are innervated by multiple motor neurons. They stimulate motor neurons with simple bipolar metal electrodes poked through the shell of an isolated crayfish walking leg or cheliped. Dactyl closer muscle contractions are monitored by a transducer writing on a smoked drum! (Today, of course, the transducer would feed into an oscilloscope or computer with A/D board). Students define nerve stimulation regimens that facilitate muscle tension, control the rate of contraction, and demonstrate the “ratchet” effect, whereby a single extra stimulus pulse added to ongoing repetitive stimuli causes a partial contraction that increases muscle tension. Another exercise examines excitation and inhibition of crayfish claw contraction with the same crude stimulation technique. Again measuring muscle contraction, students adjust stimulus parameters to recruit fast and slow muscle fibers and determine different thresholds for recruitment of excitatory and inhibitory innervation. Next, the motor nerve is split into finer bundles and movements of the leg tip are observed with stimulation of different groups of nerve fibers. Properties of twitch, tetanus, contraction rates, relaxation, and fatigue, and the synaptic sites of these properties, are examined by comparing contractions evoked by direct muscle stimulation with those evoked by nerve stimulation. The effects of temperature and insecticides on neurally evoked contractions are suggested as ways to examine environmental modulation of the NMJ. The second edition (Welsh and Smith 1960) suggests adding the inhibitory neurotransmitter gammaaminobutyric acid (GABA) to examine its direct effects on claw muscle contraction with and without the GABA receptor blocker picrotoxin. Hoyle (1968) describes a similar exercise with the crayfish claw opener muscle in more useful detail, with background information and guided interpretation, and also describes recording of muscle EMG activity (see also Welsh et al. 1968). In addition, Hoyle (1968) describes an intracellular recording exercise with the dactyl closer muscle to examine postsynaptic potential (PSP) facilitation. These basic exercises measuring muscle contraction are important because they introduce students to principles of motor control different from vertebrates. They also set the stage for later exercises that use the polyneuronal, multiterminal, and excitatory/inhibitory innervation of the crustacean muscle as a model system for vertebrate brain synapses (Wyttenbach et al. 1999). The intracellular recording exercises of Hoyle (1968) and Atwood and Parnas (1968; see below) start a new era in student laboratory physiology. The crustacean NMJ is now a model system for the study of fundamental properties of synaptic transmission and integration. The classic leg and dactyl muscle preparation (Fig. 20.2) was recently redescribed by Cooper and colleagues in video articles with extensive background and literature review. Dissection of the opener muscle preparation is described in detail, with a demonstration of short-term synaptic plasticity recorded intracellularly from the muscle. Extracellular “macro patch” recordings measure evoked and miniature PSPs and analyze the quantal content of transmitter release after



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Fig. 20.2. Crayfish leg muscle preparations. (A) Motor nerve for the leg extensor muscle, stained with the fluorescent vital dye 4-Di-2-ASP. Examples of tonic (varicosities) and phasic (thin, less prominent synapses) neurons are labeled. (B) Diagram of dactyl opener and extensor muscle preparations. (C) Mixed tonic and phasic (upper trace) and tonic-only (lower trace) excitatory postsynaptic potentials (EPSPs) from leg extensor muscle fibers during stimulation of tonic and phasic motor neurons. Tonic EPSPs show facilitation, while phasic EPSPs are of greater amplitude. A and C adapted from Wu and Cooper (2010), with permission from MyJoVE Corporation.

visualization of single synaptic boutons with the fluorescent vital dye 4-Di-2-Asp (Cooper and Cooper 2009). The crayfish leg extensor muscle preparation is used to demonstrate high- and low-output nerve terminals that synapse on the same muscle fibers (Fig. 20.2A,B) but have very different synaptic output and short-term dynamics. Differential stimulation of motor axons shows different intracellularly recorded excitatory postsynaptic potential (EPSP) amplitudes with different short-term dynamics in the same muscle fiber (Fig. 20.2C), while extracellular macro patch recording demonstrates differing quantal content of high and low output synaptic junctions. Again, the general innervation pattern of muscle is visualized with methylene blue staining before physiology and the distribution of single synaptic boutons is observed with the fluorescent vital dye 4-Di-2-Asp (Fig. 20.2A; Wu and Cooper 2010). The American/Canadian section of the International Brain Research Organization (IBRO) also has a good description of the dactyl opener muscle preparation on its website (IBRO 2011). Abdominal Extensor Another early student exercise for recording intracellularly from the crustacean NMJ appears in Atwood and Parnas (1968). They use methylene blue to study innervation patterns of the muscle and, after formalin fixation, compare sarcomere lengths in deep and superficial extensors. Intracellular recordings from muscle fibers and stimulation of motor nerves demonstrate

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Bruce R. Johnson, Robert A. Wyttenbach, and Ronald R. Hoy facilitation and posttetantic potentiation. The effects of GABA and strychnine on PSP amplitude and the ability of picrotoxin to block the effect of GABA demonstrate inhibitory input to the muscle. Baierlein et al. (2011) provide an updated video version of the original lab exercise of Atwood and Parnas (1968). Cleland (2005) uses this preparation to pose the experimental question of transmitter identity. Students first apply toxins acting on different receptor types and note their effects on EPSP amplitude, muscle resting potential, and input resistance. Their results are correlated with the effects of these drugs pressure injected or iontophoresed onto the muscle surface during intracellular muscle recording. Abdominal Superficial Flexor Several exercises in the Crawdad CD (Wyttenbach et al. 1999, Johnson et al. 2002) are based on the crayfish abdominal superficial flexor muscle (Fig. 20.3A) and its innervation. The student manual includes detailed video of dissections and recording methods, while the instructor’s manual adds detailed setup instructions and sample results. The superficial branch of abdominal nerve 3 is purely motor and innervates the superficial flexor muscles in each segment (Fig. 20.3B). A series of four exercises is available. 1. Extracellularly record spontaneous activity of nerve 3, sort APs by amplitude or energy density (arising from motor axons of differing diameters), determine the number of neurons innervating the muscle, and characterize activity patterns of each motor neuron. Tactile stimulation of the tail fan and pleopods demonstrate reflexes that differentially alter spontaneous activity in the motor neurons (Fig. 20.3C). Baierlein et al. (2011) describe a similar exercise that characterizes nerve 3 and synaptic activity in response to sensory stimuli. 2. Backfill nerve 3 with cobalt to observe functional morphology and ganglionic location of the motor neurons. 3. Match extracellularly recorded APs in the nerve with intracellularly recorded PSPs in the muscle, finding examples of selective polyneuronal innervation of muscle fibers and examining synaptic integration (summation and excitatory/inhibitory interactions), especially during sensory stimulation (Fig. 20.4A). 4. Stimulate the nerve with pulse pairs and trains to elicit facilitation, depression, and posttetanic potentiation (Fig. 20.4B), and investigate modulation of synaptic transmission, particularly by amines. The two exercises on synaptic transmission are also described in Paul et al. (1997), who adapted our Cornell laboratory class handouts for their use.

Ionic Basis of the Resting Potential and Neuronal Excitability The large fibers of crustacean claw, leg, and tail muscles are excellent preparations in which to determine the ionic basis of the resting potential. Using intracellular recording to examine the role of K+ concentration in setting the resting potential of fibers in crayfish abdominal extensor muscle (Atwood and Parnas 1968, Baierlein et al. 2011) and superficial flexor muscle (Wyttenbach et al. 1999) has been described. Wyttenbach et al. (1999) also ask students to test the contributions of Na+, Cl–, and Ca 2+. In a more difficult preparation, students test the importance of K+ and Na+ for the resting potential of giant axons in the lobster nerve cord (Hoekman and Dettbarn 1972). Welsh and Smith (1960) briefly present two exercises that examine axonal responses to nerve stimulation. Isolated crab or crayfish leg nerves are stimulated and the resulting AP activity



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Fig. 20.3. The crayfish superficial flexor preparation. (A) Ventral view of a crayfish and cross section (at the indicated line) showing the main muscle groups. (B)  Ventral view of a crayfish tail segment with cuticle removed, stained with methylene blue. Ventral nerve cord, vnc; third and fourth segment ganglia (g3, g4); ganglionic nerves (n1, n2, n3); superficial flexor muscle (sf) and its attachment point (ma); pleopod stumps (pl); sternites cut along midline (s3, s4). (C) Extracellular recording of motor neuron action potentials from nerve 3. Top trace, activity after telson stimulation; middle trace, activity after pleopod (swimmeret) stimulation; bottom trace, spontaneous activity without stimulation. Distinct action potential amplitudes indicate firing from individual axons of different diameters. Adapted from Wyttenbach et al. (1999), with permission from Sinauer Associates, Inc.

recorded extracellularly. Reduced or calcium-free saline and DDT are tested for their ability to induce repetitive firing in sensory and motor neurons. In the third edition (Welsh et al. 1968), students examine the effect of stimulus strength and duration on nerve activity and repetitive AP firing after long-duration stimuli. Absolute and relative refractory periods of compound APs and the maximum AP firing rate are determined. Students also explain the shape of recorded nerve activity after changing the distance between bipolar recording electrodes or crushing the nerve between electrodes. A similar but more detailed exercise described by Oakley and Schafer (1978) uses Limulus leg nerves, but they suggest that lobster or crayfish leg nerves can substitute. The lobster giant axons are used to examine the effects of membrane potential, external Ca 2+ and Mg2+, acetylcholine and some of its analogs, and procaine on stimulated AP initiation, amplitude, and waveform (Hoekman and Dettbarn 1972). This detailed study of excitability is best suited to advanced students. Spontaneous activity in semi-intact and isolated ventral nerve cords of crayfish is manipulated with nicotine, picrotoxin, and ethanol in Welsh and Smith (1960,1968). They suggest blowing tobacco smoke directly on the preparation to apply nicotine, not an acceptable delivery method in today’s laboratory! (We would more likely purchase acetylcholine or other nicotinic receptor agonists from a chemical supplier). Students compare the relative

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Fig. 20.4. Spontaneous and evoked activity at the superficial flexor neuromuscular junction. (A) Top: Recording setup for matching presynaptic action potentials with muscle EPSPs. Bottom: Traces show selective and polyneuronal innervation of the muscle fiber and summation of synaptic potentials. (B) Top: Recording setup for evoking and recording EPSPs in response to paired stimulation of nerve 3 stimulation at varying intervals. Bottom: Dependence of synaptic facilitation strength on interval, showing student data fit with an exponential equation. Inset: single trace showing calculation of the facilitation index as the ratio of the amplitudes of two EPSPs elicited by paired 20-msec stimulations of the same motor neuron. Adapted from Wyttenbach et al. (1999), adapted with permission from Sinauer Associates.

occurrence of small, medium, and large fibers in cross sections of the ventral nerve commissures with the representation of small, medium, and large APs in their recordings of spontaneous activity. Welsh et  al. (1968) suggest determining AP conduction velocities by measuring travel time between two recording points a known distance apart in decapod leg and ventral nerve cords. Students compare preparations with different axonal diameters (leg nerves vs. ventral nerve cord) and under different recording conditions of external resistance. Robinson et al. (2011) update this exercise using the crayfish ventral nerve cord to show axon recruitment with increasing stimulus intensity and measure conduction velocity of the compound AP. Students also quantify absolute and relative refractory periods with twin pulse stimuli. The electrical synapse blocker heptanol is used to modify the waveform and velocity of the compound AP, showing the contribution of gap junctions to AP transmission between giant axons in the ventral nerve cord. Temperature changes can also affect the waveform and velocity of compound APs. Johnson et al. (2009) describe simultaneous measurement of conduction velocities of up to six motor neurons firing tonically in the branch of abdominal nerve 3 that innervates the superficial f lexor muscle, while varying temperature (Fig. 20.5). The nerve is cut near its entrance into the muscle and stretched across the muscle to allow the longest distance between extracellular recording points. An advantage of this preparation over the leg nerve and ventral cord is that velocities are determined for multiple identified axons of different diameters in the same recording, with no stimulation required. The disadvantage is the relatively short length over which to calculate velocity. These exercises are framed in the context of evolutionary pathways for regulating conduction velocity, such as axon diameter changes and myelination.



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Sensory Physiology The abdominal MRO of crayfish is a classic preparation with which to study sensory responses (Fig. 20.6). All published exercises using this proprioceptor investigate stimulus-response properties, different stimulus thresholds, and adaptation time courses of the slowly and rapidly adapting receptors. The preparation is described in limited to moderate detail in Welsh and Smith (1960) and Welsh et al. (1968). More development is found in the Crawdad CD (Wyttenbach et al. 1999, Johnson et al. 2002) and the video article by Leksrisawat et al. (2010). These descriptions include methylene blue staining for students to view the anatomy of the stretch receptors and suggest further exploration of neuromodulatory actions on MRO responses. The propodite-dactylus organ of the decapod leg is another proprioceptor suggested for lab exercises. This organ is one of the series of joint receptors that monitor limb segment positions. It spans the joint between the last two leg segments and is composed of tens of bipolar sensory neurons sensitive to joint movement. Descriptions of this preparation by Welsh and Smith (1960) and Welsh et al. (1968) are very brief. Methylene blue staining is used to visualize the organ, and extracellular recording during dactyl movement shows the response pattern, directional sensitivity, and adaptation properties. The crayfish caudal photoreceptor is described as a student exercise by Welsh and Smith (1960) and Welsh et  al. (1968). Unlike most preparations, which are best studied shortly after dissection, the authors suggest maintaining ventral nerve cords 12–24 h before student

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Fig. 20.6. Stimulus-response properties of the slowly adapting stretch receptor (MRO1) in crayfish. (A) Diagram showing how flexing the tail stretches the superficial extensor muscle and stimulates the stretch receptor. (B) Extracellular recording of MRO1 response to a maintained stretch, showing adaptation of action potential firing. (C) Stimulus-response curve for MRO1. (D) Instantaneous spike rate of MRO1 plotted against time for maintained stretches of different amounts. Adaptation curve for each stretch fit with an exponential equation (time constant ~ 4 sec). Adapted from Wyttenbach et al. (1999), with permission from Sinauer Associates.

experiments in order to reduce spontaneous activity of other cord neurons. Extracellular recording from the ventral nerve cord with focused illumination shows the location of light-sensitive interneurons, the latent period and after-discharge of the photoreceptor after illumination, the stimulus-response relationship, and the flicker-fusion response to repeated stimuli. In a much more detailed exercise, Hermann (1972) asks for a rigorous mathematical analysis of the caudal photoreceptor response. The students also examine the receptor’s response after applying GABA and acetylcholine. The aim of this exercise is to determine the features of the receptor signal important to initiate a behavioral response such as walking. Summed photoreceptor responses in eyes (electroretinograms, or ERGs) are easily recorded in many arthropods, including crustaceans. Olivo (2003) briefly describes recording crayfish ERGs with pin electrodes, with full details on his course website (Olivo 2012). This exercise focuses on the role of second messengers in visual transduction, and students quantify stimulus-response relationships. A simple wick electrode placed on the crayfish eye, adapted from a lab exercise using Limulus eyes (Wald et  al. 1962), would also record light responses (Olivo, personal communication). Central Neural Networks The neural network generating rhythmic movements of pleopods in the crayfish tail is an excellent preparation that has not received the attention it deserves for teaching. It allows examination of a central pattern generating network, leading to comparisons and discussion of the



Crustaceans as Model Systems for Teaching Neuroscience

mechanisms organizing repetitive rhythmic movement, such as locomotion, in all animals. The best description for a student exercise is on a course website (Olivo 2012) with a brief published description in Olivo (2003). Welsh et  al. (1968) very briefly describe an exercise to characterize sensory interneurons in the crayfish ventral nerve cord and circumesophageal connectives. Students are directed to determine the response properties, location of receptive modalities and fields, and direction of signal travel in interneurons responding to tactile or visual stimuli. Although not presented specifically as a student exercise, Herberholz (2009) describes a simple measure of central network activity that could easily be done by students. Wire recording and ground electrodes are positioned in a small water-filled chamber to record neural and muscular activity during escape responses elicited by disturbing a crayfish. This is combined with behavioral observations and/or high-speed video recording for later correlation with nerve and muscle activity. Students can record activity from neural networks that mediate strong escape responses to head and tail stimuli, and a weaker avoidance response to milder touch of the thorax. Results are discussed in the context of the organization of neural networks underlying each behavior and the social conditions that may alter the efficacy of the networks. Control of Heart and Respiratory Function Crustaceans have been popular teaching models for cardiac and, to a lesser extent, respiratory physiology. The first edition of the Welsh and Smith series (1949) has several such exercises. The cladoceran Daphnia is used to determine the sensitivity of heart and respiration rates to temperature changes through direct observation and calculation of the Q10 (Welsh and Smith 1949, 1960, Welsh et al. 1968). A version of this exercise has also been developed for high school students to examine the pharmacology of heart function (Biotechnology Institute 2002). However, most exercises examining heart function and its neural control use decapods. Welsh and Smith (1949) describe an exercise to monitor spontaneous heart rate in response to application of acetylcholine and epinephrine in the isolated heart. Students compare the effects of these transmitters on neurogenic crustacean hearts with their effects on the myogenic vertebrate hearts. In the 1960 edition, Welsh and Smith added serotonin and pericardial organ extract from crabs to be tested as potential hormonal controls of heart rate. Florey (1968a) describes the crustacean heart in more detail and a semi-intact preparation to study control of the crayfish heart. Since crustacean hearts are neurogenic, heartbeat frequency reflects the burst frequency of the motor neurons, contraction amplitude reflects the number of APs per burst, and contraction duration reflects the burst duration (Florey 1968a). Experiments include stimulating cardiac accelerator and inhibitory nerves separately or together to determine their effects on heartbeat parameters. A separate exercise combines nerve stimulation with pharmacological study of excitatory (glutamate, acetylcholine, atropine, eserine, and epinephrine), inhibitory (GABA and picrotoxin), and possible hormonal (serotonin) control of heart rate (Florey 1968b). The goal is to test hypotheses of transmitter identity and whether the transmitters/modulators act within the neural network for heart regulation, at the heart NMJ, or both. A comparison of the pharmacology of the neurogenic heart of lobsters and the myogenic heart of clams, measuring heart rate and contraction strength during application of acetylcholine, GABA, and serotonin, is described in detail by Zamer and Shick (2005). Bierbower and Cooper’s exercise (2009) monitors heart and respiration rates in freely moving crayfish with extracellular electrocardiogram (ECG) recordings. This straightforward exercise is designed as a physiological measurement of the animal’s response to environmental and social stressors that might not be obvious with strictly behavioral observations. Although specific protocols are not given, DiCecco et al. (2007) describe a lab course for engineering students that uses methylene

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Bruce R. Johnson, Robert A. Wyttenbach, and Ronald R. Hoy blue to examine heart network anatomy and intracellular recording to examine properties of lobster heart neurons. This is one of several invertebrate preparations that DiCecco et al. (2007) used to teach biomedical engineers dissection, standard laboratory practices, electrophysiology, application of engineering concepts to neural networks, and data acquisition and analysis. In our Cornell neurophysiology lab course, we have also seen growing interest by engineering students in hands-on neurobiology work over the past few years. Rhythmicity of Intestinal Muscle The crayfish hindgut has been developed as a preparation to study the pharmacology of the enteric nervous system. Spontaneous contractions of intact (Cooper et  al. 2011)  or isolated (Florey 1968c, Cooper et al. 2011) crayfish hindguts are monitored under perfusion with various neurotransmitter candidates including acetylcholine, atropine, eserine, glutamate, GABA, picrotoxin, serotonin, and dopamine. Students observe the effects on the rate and strength of contraction, and construct dose-response curves for each effect. The Florey (1968c) exercise has detailed methods and guided exploration; the Cooper et al. (2011) exercise has excellent background information and video of the procedure. Both exercises frame the exercise in the context of transmitters acting both pre- and postsynaptically to shape motor output. Neurogenesis Paul et al. (2002) use 5-bromo-2’-deoxiuridine to label newborn cells in the crayfish brain as a student exercise to examine the importance of environmental stimulation for brain development (Fig.  20.7; see c­ hapter  7 in this volume for more on this technique). Students observe greater neurogenesis in animals given “enriched” environments (presence of many conspecifics, water aeration, frequent feedings, many objects for manipulation, and hiding places) than in those in “impoverished” environments (living with mother and siblings only, no water aeration, less frequent feeding, minimal external stimulation, no hiding places, very shallow water).

A

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Fig. 20.7. Neurogenesis in crayfish brains. Confocal images of juvenile crayfish brains from enriched (A) and impoverished (B) environments. Bright areas in A (indicated by arrow heads) demonstrate neurogenesis in a crayfish from an enriched environment. Adapted from Paul et al. 2002, with permission from the Faculty for Undergraduate Research.



Crustaceans as Model Systems for Teaching Neuroscience

Educational Simulations Although often not stated specifically, crustacean research has contributed to computational simulations that teach principles of cellular and systems neuroscience. For example, the “Swimmy” software (Grisham et al. 2008) uses fish swimming to guide students through the properties of a central pattern generator based on an understanding of these networks and their cellular components from crustacean research (see ­chapter 5 in this volume). A computer simulation tool that highlights crustaceans is AnimatLab (Cofer et al. 2010, AnimatLab 2011). This is an interesting project that could be very useful for student neuromechanical models of behavioral systems. Example models are presented for crayfish escape behavior and walking-leg movements, and there is potential for student development of other systems. Equipment and Software Our work developing crustacean neurobiology preparations for teaching neuroscience also motivated us to design inexpensive hardware specifically for the student laboratory. These ranged from build-it-yourself extracellular amplifiers (Land et al. 2001), nerve stimulation circuits (Land et al. 2004), and suction electrodes (Johnson et al. 2007), to inexpensive manipulators (Krans et al. 2006). In addition, we designed free software for data acquisition and analysis that highlights analysis of crayfish nerve 3 activity, synaptic physiology, and MRO responses (Lott et al. 2009). Commercially Sponsored Material Some suppliers of research equipment have produced lab handouts for faculty to use crustaceans in laboratory teaching. These are usually adapted to a specific manufacturer’s equipment, but are still useful as lab exercise guides. The descriptions usually have limited background and lack the depth of exploration found in published lab exercises. For example, iWorx offers crayfish exercises (iWorx 2011a), similar to those in the Crawdad project, that examine (1) variability in resting potential of abdominal fast extensor muscles and the effects of changing external concentration of Na+ and K+; (2) matching of motor nerve APs and muscle synaptic potentials in crayfish superficial flexor muscle, with motor nerve stimulation to determine thresholds for initial and maximal EPSP generation; and (3) stimulus-response properties and adaptation of abdominal MROs. In addition, their newsletter (iWorx 2011b) offers exercises for examining effects of temperature changes and pharmacological agents on ECG and muscle tension in crayfish heart, and the pharmacology of crayfish hindgut contractions. ADInstruments offers downloads of working guides designed to complement the Crawdad lab exercises using ADInstruments equipment (ADInstruments 2011a). There is also a “Peer Submitted Lab Chart Experiment” section (ADInstruments 2011b) including some exercises described above (Leksrisawat et al. 2010, Baierlein et al. 2011, Cooper et al. 2011, Robinson et al. 2011), and an exercise examining heart rate in the transparent shrimp Palaemonetes kadiakensis in response to temperature changes and nicotine. Although not designed for teaching, the handouts produced by the Grass Instrument Company (now Grass Technologies, a subsidiary of Astro-Med, Inc.) are educational. They describe live neurophysiological demonstrations at annual meetings of the Society for Neuroscience between 1969 and 1993. These delightful handouts highlight a variety of preparations including five dedicated to crustaceans: (1) ECG recording from the American lobster and the effect of temperature on heart and respiration rates (1973); (2) crayfish ERG recording, with brief descriptions of the abdominal stretch receptor and crayfish escape response (1976); (3) claw

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Bruce R. Johnson, Robert A. Wyttenbach, and Ronald R. Hoy asymmetry in fiddler crabs using EMG recordings from the major claw, and ECG recordings from freely moving animals (1982); (4) conduction velocity of axons in the crayfish ventral nerve cord (1985); and (5) responses of crayfish abdominal MROs (1990). These handouts were often prepared with the help of crustacean neurobiologists. The methods are not detailed enough for student directions, but the background information is often rich with extensive bibliographies. They are a good guide for an experienced neurophysiologist to develop a class exercise. The full set of handouts is now available online (Grass Technologies 2011).

BEHAVIORAL EXERCISES WITH CRUSTACEANS We draw attention to a few behavioral lab exercises because they easily lead to discussion of neural mechanisms. For example, the older Harvard lab manuals have brief descriptions of behavioral exercises that could be further developed in a neuroethological context. These include photo- and geotaxis in Daphnia; social, environmental, and physiological conditions controlling leg autotomy in crabs; control of eye and leg posture by statocyst balance organs in crabs; visual acuity and flicker fusion responses to rotating striped drums in crabs; and the measurement of diurnal rhythms in crabs (Welsh and Smith 1949, 1960, Welsh et al. 1968). Environmental control of pigment dispersion for body coloration in shrimp is described by O’Halloran (1990; see also Oakley and Schafer 1978); students consider whether body coloration is under humoral or neural control. To examine hormonal control of antagonistic behavior, Mead (2008) describes a lab exercise for non-biology majors, with an evaluative component, in which students compare male crayfish fighting behavior between controls, sham-operated animals, and animals with their androgenic glands removed. Students favorably reported that the exercise was interesting, they learned to quantify behavior, they better understood the link between hormones and aggression, and they were motivated to do more science. An exercise examining the visual contribution to dominance hierarchy establishment is described in a video article by Mercier and May (2010). Crayfish responses to their own reflections and to other crayfish in adjacent tanks are observed and discussed in the context of the sensory control of social behavior.

INSECTS AND OTHER ARTHROPODS We mention a small selection of insect neurobiological exercises because these also demonstrate basic principles of nervous system physiology in small nervous systems. Insects and crustaceans can often substitute for each other in student lab exercises. For example, an exercise in Welsh et al. (1968) examines chemosensory hair stimulation in insects and suggests that crayfish can be substituted. The classic student preparation examining coding of stimulus intensity and sensory adaptation in the cockroach leg appears in the early Harvard lab manuals. Updated versions of this exercise are found in Oakley and Schafer (1978) and Linder and Palka (1992), and a simplified version of this exercise is being developed for high school students (Marzullo and Gage 2012). Descriptions of responses of cockroach interneurons to cercal sensory stimulation recorded from the ventral cord are found in Welsh and Smith (1949, 1960), Welsh et al. (1968), and Oakley and Schafer (1978). A series of linked lab exercises describe recordings of sensory stimulation of the cockroach leg and antennae, and sensory interneuronal activity in the ventral nerve cord (Ramos et al. 2007). Other recent sensory lab exercises using insects examine the physiology of taste reception in flies (Pollack 2005a), central processing of wind information in crickets (Pollack 2005b), ERG recordings from eyes of flies (Krans et al. 2006), and proprioceptive coding of wing movements in flying locusts (Gray and Robertson 2005). The generation



Crustaceans as Model Systems for Teaching Neuroscience

of rhythmic motor patterns is examined in locust flight (Dawson and Meldrum 2005; see also Welsh and Smith 1949) and the tobacco hornworm (Trimmer 2005). Articles describing experimental protocols on synaptic physiology of the NMJ of Drosophila (Zhang and Stewart 2010) and recording of the giant fiber system of flies (Allen and Godenschwege 2010) facilitate the use of these classic preparations for the student teaching laboratory. Finally, recent advances in optogenetics have made fruitfly preparations more practical for the undergraduate student teaching lab by controlling neural activity through temperature and light stimuli (Berni et al. 2010, Pulver et al. 2011).

FUTURE DIRECTIONS Many of the crustacean teaching preparations described in early lab manuals could be further developed and updated for the student neuroscience laboratory. For example, neural control of rhythmicity of the heart and hindgut could be more widely used models for efferent control of rhythmic activity, while reconnecting crustacean neuromuscular physiology with claw movement and/or muscle tension would emphasize the functional importance of motor innervation strategies and synaptic plasticity in crustaceans and insects. Further development of the crayfish ventral photoreceptor, ERG, and leg proprioception will provide alternative models for sensory physiology. The pleopod neural network is one of the most accessible models for study of central pattern generation and could be intellectually packaged to reach a broad audience of students. Refining these older exercises presents more modern options for demonstrating core principles of signal transmission in the nervous system. Exercises on coordination and interaction of segmental motor networks and on sensory processing could introduce students to systems neuroscience and complex integration by neural networks. Examples are coordination of nerve 3 postural motor activity and central pattern generator activity for pleopod movement (see ­chapter 5 in this volume) across crayfish tail segments, and visual integration in crayfish optic ganglia (Glantz and Miller 2002; see c­ hapter 8 in this volume). Almost all published lab exercises using crustaceans do not have student evaluative components (but see Mead 2008 above). Future development of teaching exercises should include an assessment of their educational effectiveness. As other disciplines, such as engineering and computational biology, continue to take advantage of small systems neurobiology for teaching, new teaching synergies will develop. For example, DiCecco et al. (2007) describe the “Cricket Car,” made by engineering students, which translates cricket EMG recordings related to ultrasound/collision avoidance into steering directions for a remote-controlled car. In addition, the AnimatLab project (Cofer et al. 2010) combines small systems neurobiology with computational strategies for simulations of neuromechanical models. Finally, molecular analysis techniques applied to important nervous system molecules, such as amine receptors in crustaceans, are now transferable to student laboratories (McCoole et al. 2012; Christie, pers. comm.). Bioinformatics-based exercises (Grisham 2009) could also take advantage of advances in genomics and proteomics to inspire the development of crustacean teaching exercises for the 21st century.

CONCLUSIONS Crustacean neurobiology has contributed immensely to the teaching laboratory toolbox for hands-on learning of basic principles of nervous system physiology. A relatively large selection of crustacean teaching preparations were represented in early published lab manuals. Presently,

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Bruce R. Johnson, Robert A. Wyttenbach, and Ronald R. Hoy however, it appears that only a few preparations are used widely for studies of resting potential, synaptic transmission, and sensory physiology. Many older preparations not commonly in use could be modernized for the millennial student, and there are yet more crustacean research preparations that could be developed for general student studies of nervous system principles. The impact of crustacean neurobiology on the understanding of nervous system physiology will grow as students and faculty from varied disciplines apply their specific tools and paradigms to neuroscience questions.

ACKNOWLEDGMENTS Our teaching developments were supported by NSF award 9555095, an HHMI Professor award to RRH, and the Department of Neurobiology and Behavior, Cornell University. We thank our teachers, mentors, and colleagues who taught us crustacean neurobiology including Don Kennedy, Barry Ache, Jelle Atema, Mike Mellon, Fred Lang, C.K. Govind, Ed Kravitz, Ron Harris-Warrick, and Eve Marder, and our colleagues who taught and/or helped develop crustacean neurobiology labs with us, including Mike May, Peter Brodfuehrer, Gus Lott, Steve Hauptman, Farzan Nadim, Jorge Golowash, Dawn Blitz, and Wolfgang Stein. We also thank Tina Pollard of Astro-Med, Inc., for providing us with the handouts from Grass Instrument Company and making them now easily available online (Grass Technologies 2011).

REFERENCES ADInstruments. 2011a. Neurophysiology—Education—ADInstruments. http://www.adinstruments. com/solutions/education/neurophysiology/ ADInstruments. 2011b. Peer-Submitted LabChart Experiments—ADInstruments. http://www. adinstruments.com/support/submitexperiments/labchart/ Allen, M.J., and T.A. Godenschwege. 2010. Electrophysiological recordings from the Drosophila giant fiber system. Pages 215–224 in B. Zhang, M.R. Freeman, and S. Waddell, editors. Drosophila neurobiology: a laboratory manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. AnimatLab. 2011. Neuromechanical and Biomechanical Simulation. http://www.animatlab.com/ Atwood, H.L. 2008. Parallel ‘phasic’ and ‘tonic’ motor systems of crayfish abdomen. Journal of Experimental Biology 211:2193–2195. Atwood, H.L., and I. Parnas. 1968. Recording from the crayfish abdominal extensor muscle preparation with microelectrodes. Pages 307–330 in G.A. Kerkut, editor. Experiments in physiology and biochemistry, Vol. 1. Academic Press, London. Baierlein, B., A.L. Thurow, H.L. Atwood, and R.L. Cooper. 2011. Membrane potentials, synaptic responses, neuronal circuitry, neuromodulation and muscle histology using the crayfish: student laboratory exercises. Journal of Visualized Experiments 47:e2322, doi: 10.3791/2322. Berni, J., A.M. Muldal, and S.R. Pulver. 2010. Using neurogenetics and warmth-gated ion channel TRPA1 to study the neural basis of behavior in Drosophila. Journal of Undergraduate Neuroscience Education 9:A5–A14. Bierbower, S.M., and R.L Cooper. 2009. Measures of heart and ventilatory rates in freely moving crayfish. Journal of Visualized Experiments 32:e1594, doi: 10.3791/1594. Biotechnology Institute. 2002. Something you can try: the tell-tale heart. http://www.biotechinstitute. org/node/1276 Cleland, T.A. 2005. Pharmacology of the crayfish neuromuscular junction. Pages 107–127 in D.U. Silverthorn, B.R. Johnson, and A.C. Mills, editors. Laboratory manual for physiology. Pearson Education, San Francisco.



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Cofer, D., G. Cymbalyuk, J. Reid, Y. Shu, W.J. Heitler, and D.H. Edwards. 2010. AnimatLab: a 3D graphics program for neuromechanical simulations. Journal of Neuroscience Methods 187:280–288. Cooper, A.S., and R.L. Cooper. 2009. Historical view and demonstration of physiology at the NMJ at the crayfish opener muscle. Journal of Visualized Experiments 33: e1595, doi: 10.3791/1595. Cooper, A.S., B. Leksrisawat, A.J. Mercier, A.B. Gilberts, and R.L. Cooper. 2011. Physiological experimentations with the crayfish hindgut. Journal of Visualized Experiments 47:e2324, doi: 10.3791/2324. Dawson, J.W., and R.R. Meldrum. 2005. Motor patterning: electromyographic recording from wing muscles during flight in the locust. Pages 129–145 in D.U. Silverthorn, B.R. Johnson, and A.C. Mills, editors. Laboratory manual for physiology. Pearson Education, San Francisco. Deyrup-Olsen, I., and T.M. Linder. 1991. Use of invertebrate animals to teach physiological principles. Advances in Physiology Education 260:S22–S24. DiCecco, J., J. Wu, K. Kuwasawa, and Y. Sun. 2007. A novel approach to physiology for biomedical engineering students. Advances in Physiology Education 31:45–50. Florey, E. 1968a. The function of the cardioregulator nerves in the crayfish heart. Pages 246–254 in G.A. Kerkut, editor. Experiments in physiology and biochemistry, Vol. 1. Academic Press, London. Florey, E. 1968b. Pharmacology of the crayfish heart. Pages 255–260 in G.A. Kerkut, editor. Experiments in physiology and biochemistry, Vol. 1. Academic Press, London. Florey, E. 1968c. Spontaneous activity of the crayfish hindgut and its control by drugs. Pages 260–267 in G.A. Kerkut, editor. Experiments in physiology and biochemistry, Vol. 1. Academic Press, London. Florey, E. 1990. Crustacean neurobiology: history and perspectives. Pages 4–32 in K. Wiese, W.-D. Krenz, J. Tautz, H. Reichert, and B. Mulloney, editors. Frontiers in crustacean neurobiology. Birkhäuser Verlag, Basel. Glantz, R.M., and C.S. Miller. 2002. Signal processing in the crayfish optic lobe: contrast, motion and polarization vision. Pages 486–498 in K. Wiese, editor. The crustacean nervous system. Springer-Verlag, Berlin Heidelberg. Grass Technologies. 2011. Grass Technologies Knowledgebase—Application Notes. http://www. grasstechnologies.com/knowledgebase/appnotes.html Gray, J.R., and R.M. Robertson. 2005. Sensory coding: extracellular recording from the wing hinge stretch receptor of the locust. Pages 297–306 in D.U. Silverthorn, B.R. Johnson, and A.C. Mills, editors. Laboratory manual for physiology. Pearson Education, San Francisco. Grisham, W. 2009. Modular Digital Course in Undergraduate Neuroscience Education (MDCUNE): a website offering free digital tools for neuroscience educators. Journal of Undergraduate Neuroscience Education 8:A26–A31. Grisham, W., N.A. Schottler, and F.B. Krasne. 2008. SWIMMY: free software for teaching neurophysiology of neuronal circuits. Journal of Undergraduate Neuroscience Education 7:A1–A8. Herberholz, J. 2009. Recordings of neural circuit activation in freely behaving animals. Journal of Visualized Experiments 29:E1297, doi:10.3791/1297. Hermann, H.T. 1972. Analysis of the properties of the crayfish caudal photoreceptor (PRUphotoreceptor unit). Pages 155–192 in G.A. Kerkut, editor. Experiments in physiology and biochemistry, Vol. 3. Academic Press, London. Hoekman, T.B., and W.-D. Dettbarn. 1972. Neurophysiological experiments using single giant axons of the lobster. Pages 39–67 in G.A. Kerkut, editor. Experiments in physiology and biochemistry, Vol. 3. Academic Press, London. Hoyle, G. 1968. Peripheral inhibition in crayfish. Pages 280–287 in G.A. Kerkut, editor. Experiments in physiology and biochemistry, Vol. 1. Academic Press, London. IBRO. 2011. USCRC for IBRO—Resources. http://dels-old.nas.edu/USNC-IBRO-USCRC/resources_ methods_crayfish.shtml iWorx. 2011a. iWorx Neurobiology. http://www.iworx.com/content/?id=32 iWorx. 2011b. iWorx Newsletter. http://www.iworx.com/newsletter/

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Bruce R. Johnson, Robert A. Wyttenbach, and Ronald R. Hoy Johnson, B.R., R.A. Wyttenbach, and R.R. Hoy. 2002. The Crawdad Project: crustaceans as model systems for teaching principles of neuroscience. Pages 285–301 in K. Wiese, editor. Crustaceans as experimental systems in neurobiology. Springer-Verlag, Berlin Heidelberg. Johnson, B.R., S. Hauptman, and R. Bonow. 2007. Construction of a simple suction electrode for extracellular recording of nerve and muscle action potentials. Journal of Undergraduate Neuroscience Education 6:A21–A26. Johnson, B.R., S. Hauptman, and G.K. Lott. 2009. Simultaneous measurement of action potential velocity from multiple motor neurons with different conduction speeds in a tonically firing nerve from crayfish. Program No. 23.4. Neuroscience, Annual Meeting, Chicago, IL. Society for Neuroscience. Krans, J., C. Gilbert, and R.R. Hoy. 2006. Teaching insect retinal physiology with newly designed, inexpensive micromanipulators. Advances in Physiology Education 30:254–261. Land, B.R., R.A. Wyttenbach, and B.R. Johnson. 2001. Tools for the student physiology lab: an inexpensive high-performance amplifier and suction electrode for extracellular recording. Journal of Neuroscience Methods 106:47–55. Land, B.R., B.R. Johnson, R.A. Wyttenbach, and R.R. Hoy. 2004. Tools for physiology labs: inexpensive equipment for physiological stimulation. Journal of Undergraduate Neuroscience Education 3:A30–A35. Leksrisawat, B., A.S. Cooper, A.B. Gilberts, and R.L. Cooper. 2010. Response properties of muscle receptor organs in the crayfish abdomen: a student laboratory exercise in proprioception. Journal of Visualized Experiments 45:e2323, doi:10.3791/2323. Linder, T.M., and J. Palka. 1992. A student apparatus for recording action potentials in cockroach legs. Advances in Physiology Education 262:S18–S22. Lott, G.K. III, B.R. Johnson, R.H. Bonow, B.R. Land, and R.R. Hoy. 2009. g-PRIME: a free, windows based data acquisition and event analysis software package for physiology in classrooms and research labs. Journal of Undergraduate Neuroscience Education 8:A50–A54. Marzullo, T.C., G.J. Gage. 2012. The SpikerBox: a low cost, open-source bioamplifier for increasing public participation in neuroscience inquiry. PLoS ONE 7(3):e30837. Mead, K.S. 2008. Crayfish aggression and the androgenic gland in a behavior lab for nonmajors. Journal of Undergraduate Neuroscience Education 6:A60–A63. McCoole, M.D., N.J. Atkinson, D.I. Graham, E.B. Grasser, A.L. Joselow, N.M. McCall, A.M. Welker, E.J. Wilsterman Jr., K.N. Baer, A.R. Tilden, and A.E. Christie. 2012. Genomic analyses of aminergic signaling systems (dopamine, octopamine and serotonin) in Daphnia pulex. Comparative Biochemistry and Physiology 7D: 35–58. Mercier, A.J., and H.Y. May. 2010. Recording behavioral responses to reflection in crayfish. Journal of Visualized Experiments 39:e1956, doi: 10.3791/1956. O’Halloran, M.-J. 1990. Color control in shrimp. Pages 15–26 in C.A. Goldman, editor. Tested studies for laboratory teaching, Vol. 11. Association for Biology Laboratory Education (ABLE), Springfield, MO. Oakley, B., and R. Schafer. 1978. Experimental neurobiology. University of Michigan Press, Ann Arbor. Olivo, R.F. 2003. An online lab manual for neurophysiology. Journal of Undergraduate Neuroscience Education 1:A16–A22. Olivo, R.F. 2012. Bio 301 Laboratories. http://www.science.smith.edu/departments/NeuroSci/courses/ bio330/labs.html Parfitt, K. 2002. Designing an effective, affordable laboratory course in neurophysiology. Crawdad: a CD-ROM Lab manual for neurophysiology. Journal of Undergraduate Neuroscience Education 1:R5–R6. Paul, C.A., B.S. Beltz, and J. Berger-Sweeney. 1997. Discovering neurons: the experimental basis of neuroscience. Cold Spring Harbor Press, Cold Spring Harbor, New York. Paul, C.A., E.M. Goergen, and B.S. Beltz. 2002. Exploring neurogenesis in crustaceans. Journal of Undergraduate Neuroscience Education 1:A18–A22. Pollack, G.S. 2005a. The physiology of taste receptors of flies. Pages 657–665 in D.U. Silverthorn, B.R. Johnson, and A.C. Mills, editors. Laboratory manual for physiology. Pearson Education, San Francisco.



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Pollack, G.S. 2005b. Wind-sensitive interneurons of crickets: processing of sensory information. Pages 667–676 in D.U. Silverthorn, B.R. Johnson, and A.C. Mills, editors. Laboratory manual for physiology. Pearson Education, San Francisco. Pulver, S.R., N.J. Hornstein, B.R. Land, and B.R. Johnson. 2011. Optogenetics in the teaching laboratory: using channelrhodopsin-2 to study the neural basis of behavior and synaptic physiology in Drosophila. Advances in Physiology Education 35:82–91. Ramos, R.L., A. Moiseff, and J.C. Brumberg. 2007. Utility and versatility of extracellular recordings from the cockroach for neurophysiological instruction and demonstration. Journal of Undergraduate Neuroscience Education 5:A28–A34. Robinson, M.M., J.M. Martin, H.L. Atwood, and R.L. Cooper. 2011. Modeling biological membranes with circuit boards and measuring conduction velocity in axons: student laboratory exercises. Journal of Visualized Experiments 47:e2325, doi: 10.3791/2325. Trimmer, B.A. 2005. A central pattern generator in pupae of the tobacco hornworm, Manduca sexta. Pages 913–923 in D.U. Silverthorn, B.R. Johnson, and A.C. Mills, editors. Laboratory manual for physiology. Pearson Education, San Francisco. Wald, G., P. Albersheim, J. Dowling, Johns Hopkins III, and S. Lacks. 1962. Twenty-six afternoons of biology: an introductory lab manual. Addison-Wesley, Reading, MA. Welsh, J.H., and R.I. Smith. 1949. Laboratory exercises in invertebrate physiology. Burgess, Minneapolis. Welsh, J.H., and R.I. Smith. 1960. Laboratory exercises in invertebrate physiology. Burgess, Minneapolis. Welsh, J.H., R.I. Smith, and A.E. Kammer. 1968. Laboratory exercises in invertebrate physiology. Burgess, Minneapolis. Wiese, K., editor. 2002a. The crustacean nervous system. Springer, Berlin, Heidelberg. Wiese, K., editor. 2002b. Crustacean experimental systems in neurobiology. Springer, Berlin, Heidelberg. Wu, W.H., and R.L. Cooper. 2010. Physiological recordings of high and low output NMJs on the crayfish leg extensor muscle. Journal of Visualized Experiments 45:e2319, doi:10.3791/2319. Wyttenbach, R.A., B.R. Johnson, and R.R. Hoy. 1999. Crawdad: a CD-ROM lab manual for neurophysiology. Sinauer Associates, Sunderland, MA. Zamer, W.E., and J.M. Shick. 2005. Comparative pharmacology of lobster and clam hearts with an introduction to perfusion techniques. Pages 1073–1091 in D.U. Silverthorn, B.R. Johnson, and A.C. Mills, editors. Laboratory manual for physiology. Pearson Education, San Francisco. Zhang, B., and B. Stewart. 2010. Synaptic physiology of the Drosophila neuromuscular junction. Pages 171–214 in B. Zhang, M.R. Freeman, and S. Waddell, editors. Drosophila neurobiology: a laboratory manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York.

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INDEX

Abdominal ganglion, 58, 69, 74, 151, 227, 402, 403, 405–407, 441, 442, 448, 543 Abdominal stretch receptor, 66, 547 Abdominal tergite, 67 Acartia, 295, 300, 303 Accessory lobe, 11, 12, 14, 15, 17, 20, 21, 24, 29, 30, 33–38, 40–42, 54, 57, 185, 186, 277–279, 446, 447, 474 Accessory nerve, 72 Accessory strand, 371 Acetes, 54, 304, 305, 309, 310 Acetylcholine, 116, 120, 124, 149, 150, 221, 342, 344, 377, 378, 541, 544–546 Achelata, 2, 3, 13, 26, 38 Acrocalanus, 300 Acrophase, 439, 440, 443 Actometer, 514–517 Acute zone, 209, 210, 240, 497 Adult neuroblast, 54, 188, 190–192, 197, 279 Aesthetasc, 21–26, 29, 30, 50, 51, 53, 55–57, 69, 175, 181–183, 197, 198, 263, 266–274, 277–279, 281, 283, 306 Aesthetasc sensillum, 23–26, 29, 51, 56, 69, 263, 266, 267, 268, 270, 279 Aggregation, 22, 270, 279, 303 Aggression, 106, 117, 264, 285, 457–475, 548 Alima, 243 Alpheus, 467 Amine, 53, 57, 59, 106, 107, 115–123, 126–133, 135–137, 139, 151, 152, 192, 194, 219, 227, 265, 275, 337, 341, 345, 346, 377, 414, 416, 422, 440, 441, 443, 447, 458, 465, 467–475, 540, 546, 549 Amino acid, 52, 57, 66, 88, 117, 156, 248, 265, 273, 275, 345, 346, 443

Amphipoda, 2, 3, 8, 13, 59, 207–210, 214, 227, 238, 241, 242, 249, 251, 256, 266, 271–273, 294, 321–324, 333, 434, 435 Anaspidacea, 3, 13 Anaspides, 418 Anomala, 2, 3, 13 Anomura, 26, 34, 38, 55, 177, 179, 186, 187, 192, 194, 209, 211, 239, 266, 270, 271, 278, 368, 420 Antenna, 2, 12, 14–17, 19, 21–25, 27–34, 37, 38, 40, 52, 54, 64, 69, 74, 75, 86, 97, 180, 181, 185, 186, 192, 226, 267, 268, 272, 277, 299, 300, 304–308, 310, 313, 367, 373, 446, 447, 459, 460, 467, 548 Antenna 1, 12, 14–18, 25, 29, 30, 34 Antenna 2, 12, 14–17, 29, 30, 33, 33, 34, 40 Antennal flagellum, 27 Antennal lobe, 37, 38, 186, 277 Antennal nerve, 54, 268 Antennocerebral tract, 37, 38 Antennular flagellum, 51, 52, 55, 56, 68, 69 Antennule, 52, 54, 55, 67, 68, 73, 86, 181, 182, 185, 189, 265–273, 275, 277–279, 280, 281, 283, 284, 303, 311, 312, 465 Antromysis, 306, 309 Apoptosis, 176, 187, 189, 193, 195 Armadillidium, 23, 431 Artemia, 208, 218, 256, 303 Arteriole, 183, 191, 194 Astacida, 2, 3, 13, 26, 37, 38, 62, 74, 86, 266, 270, 299 Astacus, 28, 118, 363, 439, 469 Asymmetry, 210, 548 Atropine, 545, 546 Austropotamobius, 51, 52, 53, 309

555

556 Index Axon, 7, 20, 21, 24, 29, 33, 35, 37, 40, 52, 57–62, 71, 72, 86–94, 96, 97, 99, 103, 104, 105, 106, 108, 109, 121, 122, 124, 126, 132, 147–149, 151–153, 155, 158–161, 163–166, 176, 181, 183, 184, 186, 187, 197, 208, 212, 218, 219, 221–225, 227, 236, 266, 268, 274, 275, 277–279, 293, 300, 309, 311–314, 341, 342, 344, 366–368, 376, 382, 396–399, 404–407, 409, 411, 418, 419, 436–438, 441, 442, 521, 523, 524, 528, 536–543, 548 Balanus, 207, 215, 217, 308 Barnacle, 3, 8, 86, 207, 209, 212, 215, 217, 226, 228, 256, 264, 322 Basal body, 305, 306, 307, 309 Basal lamina, 307 Basal membrane, 447 Basis, 364, 365, 367, 369, 371–373, 381 Bathysquillidae, 18 Benthic, 13, 75, 252, 270, 272, 281, 293, 294, 296, 297, 299, 308, 310, 313, 314 Bestiolina, 295 Biological oscillation, 429 Birgus, 15, 22, 25, 26, 34, 39, 55 Bistability, 114, 116, 125, 131, 138 Boreomysis, 267 Brachyura, 2, 3, 13, 26, 34, 38, 63, 65, 70, 73, 74, 76, 86, 97, 177, 179, 186, 187, 192, 209, 221, 237, 239, 242, 254, 266, 270, 271, 278 Branchinecta, 303 Branchiopoda, 3, 179, 207–209, 211, 214, 218, 227, 266, 271, 434 Brood care, 31 Burrow, 18, 19, 23, 27, 28, 63, 209, 251, 321, 322, 397, 433, 484–488, 490, 491, 494, 497–502, 512, 513, 517 Burrow surveillance, 485, 494, 499–502 Burst, 28, 57, 89, 93, 106, 114, 116, 118–124, 126–134, 136, 138, 165, 166, 221, 276, 339, 341, 342, 345, 347, 368, 378, 383, 385, 398, 400, 401, 403, 432, 433, 545 Bythograea, 238, 242, 245 Calanus, 299 Callianassa, 25, 26, 38 Callinectes, 98, 99, 119, 121, 208, 251, 272, 363, 468 Cambarus, 19 Cancer, 25, 26, 103, 105, 116, 118, 129, 132, 187, 339, 340, 342, 346, 363, 443, 511 Caphyra, 265 Caprella, 214 Carapace, 19, 55, 63, 67, 182, 184, 212, 465, 488, 494, 497, 512, 513, 515, 524, 527 Carbachol, 121

Carcinus, 2, 4, 34, 72, 120, 178, 184, 186, 214, 223, 225, 226, 363, 365, 368, 435, 443, 463, 512 Cardiac ganglion (CG), 114–116, 118–121, 123, 149 Cardiac sac, 128–130, 135, 136, 338, 344 Caridea, 3, 13, 14, 29, 69, 75, 177, 242, 278, 299, 312, 469, 474, 475 Carpopodite, 28, 74, 88, 90 Carpus, 364, 365, 370, 376 Caudal ganglion, 20, 227, 406, 441, 447, 448 Cell enveloping, 22, 51, 56, 305–308 hypodermic, 309 lamina monopolar, 206, 219, 220 niche-forming (NFC), 189, 191–196, 198 parasol, 35, 36, 42 perivascular, 183, 191, 194 photoreceptive, 17 pigment, 207, 209, 237, 437, 496 retinula, 59–63, 207–215, 217, 218, 227, 237, 243, 245, 247, 250, 437, 447, 494, 496 sensory, 22, 66, 282, 305, 308, 309, 372 sheath, 55, 56, 180, 194, 308, 309 tapetal, 447 Cell body glia, 187, 189, 191 Cell cluster, 444–447, 474 Central nervous system (CNS) 2, 4, 12, 20, 33, 41, 50, 55, 58, 73, 75, 115, 117, 119, 147–152, 154, 156, 158, 159, 176–181, 183, 186, 193, 196–198, 277, 279, 280, 362, 363, 368, , 376, 386, 401, 422, 434, 443 Central pattern generator (CPG) 117, 118, 121–123, 125, 129, 132, 136, 159, 362, 378–380, 384–388, 398, 408, 409, 420, 547, 549 Centropages, 309 Cephalocarida, 3, 209, 236, 266, 270 Cephalothorax, 28, 67, 331, 398, 411, 417, 441 Chelae, 20, 27, 51, 52, 54, 67, 181, 272, 364, 459, 460, 462, 463 Chelicerata, 3 Cheliped, 31, 68, 71, 72, 268, 363, 538 Chemical badge, 23, 25 Chemoreception, distributed, 21, 263, 268, 275, 278–280, 283 Cherax, 16, 25–27, 58, 60, 64–66, 182, 183, 188, 211, 433, 442, 463, 465, 467 Chiasm, external, 219, 221, 223 Chiasm, internal, 219, 223 Chiasmata, 35, 184, 521, 522 Chordotonal organ, 28, 72–74, 116, 180, 182, 294, 305, 308, 362, 367, 368, 370, 381, 386, 401 Chorocaris, 212 Ciliary rootlet, 266, 271, 305, 308, 309 Circumesophageal connective, 4, 226, 379, 462, 463, 536, 545

Index 557 Cirolana, 210, 242, 265 Coding, 218, 226, 263, 273, 276, 282, 283, 347, 367–370, 373, 374, 376, 384, 548 Coenobita, 24–26, 39, 55, 194, 239, 271 Commissural ganglion (CoG), 17, 121, 123, 132, 159, 338–340, 344, 345 Communication, 23, 63, 64, 75, 76, 106, 115, 264, 270, 293, 294, 497 Competition, 252, 467, 468, 501 Conditioning, 41, 511–513, 518 Conduction velocity, 536, 537, 542, 543, 548 Contact sensilla, 52, 69 Cooperation, 32 Copepoda, 2, 3, 13, 52, 59, 60, 209, 211, 212, 214, 236, 242, 256, 264, 266, 267, 270, 271, 273, 280–282, 293–295, 299–304, 306–309, 311–314, 434, 511 Copilia, 212 Cornea, 20, 59, 61, 209–211 Corneal facet, 207–211, 237, 238 Coxa, 364, 365, 367–369, 371, 373, 381, 386 Coxopodite-basipodite chordotonal organ, 367 Crab coconut, 3, 21–25, 35, 41 fiddler, 17–20, 63, 105, 209, 214, 237–240, 248–251, 265, 267, 273, 280, 283, 284, 433–435, 440, 468, 484–497, 499–503, 512, 513, 548 hermit, 2, 3, 13, 34, 39, 55, 56, 61, 211, 271, 363, 418, 463, 464, 469, 470, 512 rock, 27 shore, 2, 4, 34, 186, 209, 435, 463, 468, 469, 512 spider, 13, 152, 160, 179, 295 Crangon, 214, 299 Crayfish cave, 17–20, 24, 25 freshwater, 17, 21, 22, 24, 25, 27, 49, 61, 251, 271, 431, 433 troglobitic, 28 Crystalline cone, 59, 61, 62, 207–211, 228, 241–243, 253255, 493, 496 Cue, chemical, 8, 12, 19, 32, 264, 272, 273, 279, 283, 285, 304, 328 Cumacea, 3, 13 Cuticle, 22–24, 52, 55, 56, 66–68, 74, 75, 177, 178, 182, 192–194, 212, 238, 267, 271, 298, 304, 306–308, 310, 313, 371, 372, 515, 520, 541 Cuticular sensilla, 50, 55, 67, 69, 180, 181, 293, 296, 304, 306 Cuticular stress detector, 363, 371, 372, 384 Cyclic adenosine monophosphate, 107, 117, 414 Cyclic guanosine monophosphate, 107, 121 Cyclops, 308

Dactyl, 52, 72, 153, 182, 251, 267, 364, 365, 371, 372, 384, 385, 538, 539, 543 Daphnia, 8, 208, 209, 214, 218, 228, 244, 249, 252, 270, 274, 285, , 434, 443, 474, 545, 548 Decapoda, 2, 3, 13–15, 37, 38, 177, 438 Decision-making, 6, 7, 413, 421, 469, 502 Defense, 31, 206, 223, 226, 265, 459, 462, 464, 465, 468, 484, 487 Deformation rate, 296, 297, 300, 314 Dendrite distal, 51, 74, 212, 305–310, 414 proximal, 306, 307, 309 sensory, 68, 305, 310 Dendrobranchiata, 3, 13, 312 Desert, 1, 11, 13, 17, 23 Detection, hydrodynamic, 296, 302 Detection, tactile, 303 Deutocerebral commissure, 37, 38 Deutocerebrum, 12, 14–17, 24, 29, 35–39, 54, 57, 175, 184–186, 188, 190, 192–196, 198, 224, 277, 279, 446, 447 Diacylglycerol, 117, 217 Digestion, 6 Dioptromysis, 210, 241 Diversity, phylogenetic, 2, 3 Dopamine, 106, 116, 119, 121, 126, 128–131, 135, 341, 414, 458, 469, 470, 475, 546 Dorsal cord interneuron, 410 Echinoderm, 31 Elasmobranch fish, 331 Electrical field, 49, 64, 66, 331 Electrical synapse, 133, 135, 147, 149, 154, 155, 337, 339, 340, 397, 398, 401–403, 405–407, 417420, 422, 542 Electromagnetic induction, 330, 331 Embryo, 2, 14, 16, 120, 175–181, 190, 193–198, 211, 349, 439 Emerita, 73, 368, 435 Endopodite, 28, 367, 401 Enteric nervous system, 546 Epilabidocera, 299 Epinephrine, 545 Eriphia, 90, 92, 94–96, 98, 118 Escape response, 63, 69, 88, 106, 223, 252, 255, 298– 302, 314, 397, 400, 409, 411–413, 415, 419, 461, 462, 464, 509, 511, 513–518, 524–526, 545, 547 Eserine, 545, 546 Esophageal ganglion, 4, 123, 338 Esophageal nerve, 16, 132 Esophagus, 4, 17, 280, 338 Euastacus, 27 Euchaeta, 267, 270

558 Index Eumalacostraca, 3, 11, 13, 14 Euphausia, 295 Euphausiacea, 3, 13, 14 Eureptantia, 13, 15, 21, 24, 30 Eurydice, 435 Evolution, 1, 6, 11, 13, 37, 38, 55, 74, 76, 104, 138, 196, 198, 210, 236, 237, 240, 242, 247, 248, 253, 255–257, 264, 270, 271, 273, 285, 300, 314, 396, 397, 406, 417–420, 423, 447, 473, 497, 542 Excirolana, 435 Excitatory junctional potential, 88, 90, 418 Excitatory postsynaptic potential, 148, 221, 374, 384, 401, 539 Exoskeleton, 4, 177, 198, 242, 417, 434 External medulla, 225, 441, 446, 448, 474, 521 Exteroceptor, 67, 69, 294, 313, 367 Eye compound, 14, 18, 58–64, 67, 175, 180, 181, 185, 198, 207, 209, 211, 212, 218, 235–240, 242, 249, 253, 255, 256, 441, 492, 495, 496, 501, 503 median, 14, 59 nauplius, 58–60, 206, 211, 212, 224, 256 Eyestalk, 2, 14, 19–21, 33, 35, 63, 69, 70, 86, 105, 122, 184–186, 209, 210, 223, 225, 227, 228, 240, 268, 278, 325, 328, 436–447, 474, 487, 495, 511, 513, 515, 520, 521 Eyestalk ganglion, 184–186, 268, 278, 439, 474 Feedback loop, 12, 429, 444, 445, 448 Fertilization, 22, 264 Fiber fast, 98–100, 366 slow-tonic, 366 slow-twitch, 366 Fiber bundle, 29, 30 Foregut, 86, 88, 114, 118, 122, 338 Fossil, 13, 76, 417 Funnel canal organ, 52, 268, 362, 372, 384, 385 Gaetanus, 295 Gamma-aminobutyric acid, 88, 127, 151, 219, 382, 414, 538 Gammarus, 273 Gastropoda, 463, 469, 470 Gaussia, 311 Genomic, 5, 8, 166, 197, 285, 474, 494, 549 Geomagnetic field, 32, 65, 322–324, 331 Gigantocypris, 59, 212 Globuli, 8, 14, 264 Glomeruli, 15, 24–26, 34, 36, 37, 40, 57, 186, 198, 277–279

Glutamate, 52, 54, 88, 116, 124, 134–136, 147, 149–152, 154, 161, 222, 223, 275, 283, 342, 379, 519, 545, 5446 Glyptonotus, 58 Goneplax, 433 Gonodactylidae, 18 Gonodactylus, 61, 62, 239, 246, 247, 250, 465 Gonopore, 22 Grapsus, 63, 208 Ground pattern, 11, 14, 20, 21, 24, 25, 29, 30, 37 Ground pattern, eureptantian, 21, 24 Ground pattern, malacostracan, 20, 21, 29, 30, 37 Habituation, 27, 158, 223, 282, 301, 314, 408, 416, 417, 422, 423, 502, 511, 513, 514, 519 Hemidesmosomes, 309 Hemiellipsoid body, 12, 14–16, 21, 24, 33–36, 38, 41, 42, 57, 184–186, 224, 444–448 Hemigrapsus, 248, 432, 433 Hemilepistus, 23 Hemimysis, 431 Hexapoda, 3, 38, 176 Hierarchy, 12, 22, 363, 457–459, 462, 467, 469, 475, 548 Hindgut, 537, 546, 547, 549 Holoplankton, 242, 294 Homarida, 2, 3, 13, 26, 38, 62, 74, 266, 270 Homarus, 16, 25, 26, 28, 35, 51, 52, 67, 68, 70–72, 75, 88, 95, 116, 129, 179, 207, 221, 274, 276, 284, 341, 363, 433, 434, 439, 458 Homing, 12, 31–33, 41, 49, 322, 327–329, 485, 487, 499 Hoplocarida, 59 Hutchinsoniella, 308 Hyas, 89 Hydrothermal vent, 1, 212, 238, 342, 244, 252, 254 Hyperia, 210 Idotea, 23, 107, 324 Immunoreactivity, 16, 119, 124, 376, 439, 441, 443–448, 472, 474 In vivo intracellular recording, 518 Inhibitory junctional potential, 93 Inhibitory postsynaptic potential, 93, 224, 343, 374, 402 Innervation, 67, 85, 87–93, 95, 97, 99, 102, 104, 105, 108, 119, 148, 154, 161, 279, 280, 364, 365, 368, 376, 414, 536–540, 542, 549 Inositol, 1, 4, 5-trisphosphate, 57, 100, 107, 415 Internal medulla, 225, 446, 447, 474, 521 Interneuron assistance reflex, 382, 384, 386 corollary discharge, 407 mechanosensory, 401, 403, 404 Intertidal zone, 63, 322, 435, 513, 516

Index 559 Ischiopodite, 28, 74 Ischium, 364, 365, 370–372, 381 Isopoda, 2, 3, 8, 11, 13, 15, 17, 23–25, 40, 54, 58–60, 76, 107, 120, 207, 209, 210, 214, 227, 228, 242, 265, 308, 321, 322, 324, 325, 333, 431, 434, 435, 443 Jasus, 25, 26, 124, 363 Kin recognition, 23, 24, 31 Krill, 13, 211, 245, 434 Labidocera, 212, 300 Lamina, 14–17, 21, 61, 64, 206, 208, 218, 221, 223, 227, 280, 307, 437, 438, 441, 444–448, 474, 521, 522, 524 Lamina ganglion, 208, 218–220, 446, 447, 521 Landmark guidance, 490, 491 Lateral flagellum, 23, 52, 55, 181–183, 268, 269 Lateral horn, 37, 38, 279 Lateral soma cluster, 184–186 Learning, 5–7, 12, 31, 33, 41, 76, 279, 396, 415, 416, 422, 509–515, 518–520, 524–529, 538, 549 Lens, 59, 61, 62, 207, 210–212, 236, 241, 242, 493, 496 Leptodora, 303 Leptograpsus, 207, 208, 217 Leptostraca, 3, 13 Libinia, 25, 26, 187 Ligia, 23, 54, 60, 120, 213, 214 Limulus, 215, 217, 219, 441, 541, 544 Lobster clawed, 2, 3, 31, 35, 36, 37, 86, 177, 179, 187, 192, 195, 207, 275, 363, 459, 464–467 spiny, 2, 3, 13, 17, 22, 24, 25, 28, 31–33, 35–37, 41, 52, 54, 55, 57, 65, 86, 116, 117, 122, 124, 177, 183, 187, 192, 195, 213, 221, 267–270, 275, 277, 279, 284, 321, 322, 325–333, 363, 364, 376, 418, 467, 474 squat, 2, 3, 207, 211, 418, 469 Lobula, 14–17, 35, 64, 206, 218, 219, 223–228, 521–525, 527 Lobula plate, 14, 15, 35, 64 Locomotion, 7, 12, 49, 93, 107, 108, 109, 225, 226, 265, 362–364, 366, 367, 372, 379–388, 429, 430, 432, 433, 435, 436, 438–445, 447, 448, 462, 469, 545 Lophogastrida, 3, 13 Lysiosquillidae, 18 Lysmata, 29 Macrobrachium, 117, 267, 440, 463, 472 Macrocyclops, 59, 60 Macrocypridina, 212 Magnetic field, 7, 32, 49, 65, 321–324, 326–328, 330–333, 487

Magnetite, 65, 321, 330–333 Magnetoreception, 6, 321, 330–333, 441 Malacostraca, 2, 3, 5, 12–15, 18–25, 29–31, 33, 37–40, 41, 54, 57–59, 67, 179, 198, 218, 224, 253, 254, 271, 273, 293–295, 299, 418, 423, 510, 521 Mandible, 16, 52, 54 Mandibular commissure, 16 Mandibulata, 3 Mate recognition, 273, 298, 303 Mating, 12, 17, 18, 22, 23, 27–31, 33, 49, 50, 63, 105, 272, 273, 303, 435, 443, 468, 502 Mating, burrow, 497–499 Mating, surface, 497–499 Maxilla, 16, 52, 54 Maxilliped, 18, 22, 28, 31, 52, 54, 74, 98, 268, 284, 309, 460 Maxillopoda, 2, 3 Mechanism, neural, 1, 7, 8, 206, 221, 226, 285, 294, 333, 447, 462, 473, 548 Mechanoreception, 6, 12, 17, 21, 22, 28–30, 41, 49, 51–54, 59, 66–73, 123, 159, 178, 181, 183, 268, 278, 293, 294, 297–300, 302–311, 313, 314, 331, 368 Medial flagellum, 67, 268, 269 Medial soma cluster, 184–186 Medulla, 11, 12, 14–17, 20, 21, 33–38, 41, 42, 57, 184, 185, 206, 214, 218, 219, 221–225, 278, 279, 441–448, 474, 521, 522, 524, 527 Memory, 5–12, 25, 31, 33, 76, 225, 279, 509–520, 522, 524–528 Meropodite, 28, 74, 153 Merus, 364, 365, 370, 376 Methylene blue, 2, 4, 185, 191, 192, 538, 539, 541, 543 Microtubules, 2, 51, 52, 54, 55, 305–308, 310, 314 Microvilli, 58, 59, 61–63, 192, 207, 208, 212–214, 251, 494, 496 Migration, seasonal, 12, 31, 333 Migration, vertical, 211, 236, 244, 255, 256, 434, 435, 443 Mollusc, cephalopod, 30, 214, 217 Molt, 2, 23, 69, 175–179, 181, 183, 198, 270, 272, 417, 429, 430, 431, 435, 464, 465 Motion detection, 206, 216, 221, 223, 224 Motor innervation, 89, 91, 279, 549 Motor nerve, 2, 20, 72, 88, 90, 107, 339, 369–371, 375, 378, 380, 383, 536, 538, 539, 547 depressor, 371 levator, 371 remotor, 375, 380 Motor neuron excitatory, 87, 88, 90, 95, 107, 109, 339 fast extensor, 376, 407, 410 fast flexor, 73, 399, 405 Movement, gastric, 5

560 Index Mudflat, 17, 63, 239, 485, 486, 492, 494, 497, 499, 501 Muscle bender, 93 closer, 88, 90, 91, 94, 96–98, 103, 105–107, 162–164, 378, 538 extensor, 73, 88, 101, 102, 104, 107, 161, 162, 378, 407, 408, 417, 536, 539, 540, 547 flexor, 72, 74, 88, 107, 365, 370, 371, 373, 378, 398, 405, 406, 408 levator, 99, 364, 378 opener, 89, 90, 148–153, 155, 156, 159, 364–366, 378, 536, 538, 539 receptor, 72, 368, 369 remotor, 97, 364, 365, 378 stretcher, 89, 90, 93, 95, 103, 105, 152, 160, 365, 539 superficial extensor, 70, 408, 539, 544 superficial flexor, 75, 149, 159, 161, 536, 540–542, 547 Muscle receptor organ, 67, 70, 313, 363, 367, 368, 381, 408, 536 Muscle tension, 106, 368, 537, 538, 547, 549 Myelination, 7, 293, 300, 311–314, 542 Myochordotonal organ, 74, 362, 368, 370 Myodocopida, 212 Myofibrillar adenosine triphosphatase, 98 Myriapoda, 3, 176 Mysidacea, 2, 3 Mysidium, 208 Mystacocarida, 3, 236 Navigation, 5, 49, 214, 235, 282, 283, 322, 327–329, 484, 485, 487, 497, 502, 503 Nekton, 59, 294 Nematoscelis, 238 Neohelice, 64, 222, 487, 509–511, 513–522, 524, 525, 528 Neomysis, 54, 253, 310 Nephrops, 100, 433 Nerve dorsal gastric, 339 motor, 2, 20, 72, 88, 90, 107, 339, 369–371, 375, 378, 380, 383, 536, 538, 539, 547 stomatogastric, 122, 346 ventricular, 116, 126, 339 Nerve cord, ventral, 2, 4, 58, 59, 70, 74, 277, 378, 379, 397, 398, 441, 442, 448, 536, 541–545, 548 Nerve impulse, 86, 96, 107, 311–313, 396 Nerve root, 4, 74, 401, 402 Neural network, 1, 114, 115, 118, 122, 125, 130, 131, 136, 138, 139, 362, 363, 372, 379, 387, 388, 449, 544, 546, 549 Neurogenesis, 6, 7, 49, 57, 76, 175–184, 186, 189, 192–199, 279, 285, 535, 537, 546

Neurogenic niche, 41, 175, 185, 186, 188–193, 197 Neuromodulation, 7, 87, 114–116, 120, 121, 131, 136–138, 343, 344, 388, 396, 422, 440, 469, 537 Neuron chemosensory, 50, 52, 54, 266, 270, 273, 276, 282–285 command, 379, 397, 400, 421, 461–463 distributed chemoreceptor, 268, 275, 278–280, 283 dorsal gastric, 127, 339 excitatory motor, 87, 88, 90, 95, 107, 109, 339 fast flexor motor, 73 gastric mill, 124, 130, 339, 343, 344, 349 inferior cardiac, 126–128, 339 inhibitory, 85, 93, 95, 104, 122 lamina monopolar, 206, 218–220 lateral giant, 73, 149, 155, 397, 400, 401, 404, 408, 417, 418, 421, 461 lobula giant, 223, 224, 487, 515, 522–525 mechanoreceptor, 51, 53, 54, 66, 73, 178, 268, 278 medial giant, 73, 155, 396, 397, 400, 418, 420, 461 non-giant, 410 olfactory receptor, 21, 24, 36, 53, 55, 56, 175, 182, 185, 268, 271, 278 phasic, 87, 94, 106 projection, 35–37, 40, 41, 57, 58, 122–125, 127, 130, 132, 137, 184, 185, 279, 343, 344, 443 pyloric, 126, 128–131, 133, 135, 347 tonic, 87–90, 94 ventral cardiac, 344 ventricular dilator, 126–128, 134, 339 Neuropeptide, 34, 108, 117, 122, 124, 125, 133, 137, 185, 187, 337, 345, 346, 438, 441, 444, 472 Neuropil antennular, 54, 57, 278, 279 brain photoreceptor, 439 cap, 35 chemosensory, 278 core, 35 glomerular, 24, 36, 37, 224 olfactory, 11 optic, 11, 18, 20, 21, 33, 35, 40, 196, 208, 214, 224, 225, 448, 520–522 primary sensory, 11, 21 protocerebral, 12, 14, 15, 36, 58, 279, 437, 445–447, 520 retinotopic, 14, 17, 218, 520 synaptic, 125, 341, 342 Neurotransmission, 14, 87–90, 93, 95, 96, 104, 106, 107, 109, 116, 117, 120, 123, 124, 137, 147, 149–152, 161, 166, 337, 345, 382, 439, 447, 473, 475, 538, 546

Index 561 Nippoleucon, 430 Norepinephrine, 116 Ocelli, 59, 212, 216, 256 Octopamine, 106, 107, 116–118, 120–123, 129, 132, 133, 135, 414, 416, 422, 458, 465, 469–474 Octopus, 30 Ocypode, 207, 209, 239, 512 Odontodactylus, 63, 215, 238, 250 Oesophageal ganglion, 123, 338 Oithona, 302 Olfaction, 21–23, 30, 39, 40, 198, 263, 268, 274 Olfaction, aerial, 22, 23, 39 Olfactory globular tract, 15, 21, 33, 35–37, 185, 224, 279 Olfactory lobe, 2, 12, 14–16, 21, 24, 25, 29, 30, 33–40, 57, 175, 184–186, 266, 268, 271, 277, 278, 446, 447, 474 Olfactory sensilla, 22, 25, 50, 55, 175, 181–183, 197 Oligostraca, 3 Ommatidia, 18–20, 59, 61–63, 175, 180, 181, 198, 206–212, 214, 218, 237–243, 247, 249–251, 437, 440, 492–494, 496, 513, 520, 521 Oplophorus, 253 Optic chiasm, 15, 521, 522 Optic ganglion, 14, 15, 19, 20, 33, 443, 444, 549 Optic lobe, 12, 16, 175, 180, 184, 185, 195, 196, 198, 206, 207, 214, 218, 219, 222, 224, 227, 228, 437, 438, 443–448, 487, 520, 522, Optical resolution, 18, 496 Orconectes, 19, 22, 71, 431, 458, 459 Pachygrapsus, 517 Pacifastacus, 121, 222, 271, 465 Pagurus, 29, 34, 56, 187, 363, 463 Palaemon, 69, 124, 253 Palaemonetes, 212, 213, 238, 252, 253, 255, 267, 431, 547 Pancarida, 3 Pancrustacea, 3 Panulirus, 24–26, 31–33, 51–53, 75, 116, 182, 183, 185, 188, 190, 191, 208, 221, 253, 267–270, 272, 309, 321, 339, 467 Parvocalanus, 300 Path integration, 6, 7, 63, 484–487, 490, 497–501 Pedomorphosis, 13 Pelagic, 240–242, 244, 249, 252–254, 256, 270, 293–299, 301–304, 309–314 Penaeus, 29, 253 Peracarida, 13, 14, 18 Percnon, 25, 26 Pereopod, 51, 52, 54, 72, 74, 98, 268, 270, 273, 299, 365

Pericardial organ, 107, 118, 119, 345, 346, 545 Periclimenes, 29 Periodicity, 429, 443 Peripheral nervous system (PNS), 176–182, 197, 198 Petrolisthes, 25, 26 Phalanx, 32 Phaso-tonic, 368, 369, 372, 374, 375 Pheromone, 22, 23, 27, 29, 264, 272, 273, 276, 278–280 Phosphatidylinositol, 4, 5-bisphosphate, 217 Phospholipase C, 107, 117, 217, 415 Photopigment, 58, 206, 213, 214 Photoreceptor, 19, 20, 40, 58–61, 63, 65, 116, 180, 181, 206, 207, 209, 212–219, 226–228, 241, 243, 245, 247–249, 254, 256, 257, 332, 428, 430, 437, 439, 440–442, 444, 447, 448, 493, 494, 496, 521, 536, 537, 543, 544, 549 Photoreceptor array, 40 Photoreceptor, brain, 58, 60, 428, 437, 439, 441, 448 Photoreceptor, caudal, 58, 59, 207, 227, 428, 440–442, 448, 536, 537, 543, 544 Phototransduction, 206, 217, 228 Phronima, 210, 238, 241 Phyllocarida, 3, 59 Phytoplankton, 214, 302, 303 Plankton, 59, 75, 211, 235, 249, 252, 255, 257, 270, 273, 280–282, 294, 434, 435 Plasticity, 7, 11, 13, 147, 148, 154, 157, 158, 163–165, 167, 196, 279, 301, 349, 388, 415, 422, 527, 537, 538, 549 Pleocyemata, 14 Pleuromamma, 52, 282, 300, 302, 306, 307, 309, 312 Polarization, 19, 60, 63, 100–103, 107, 130–132, 149, 152, 163, 206, 208, 210, 214, 215, 219–222, 225–227, 235, 239, 247, 248, 250–252, 256, 257, 347, 366, 368, 369, 373, 374, 377, 382, 402–405, 410, 415, 485, 494 Polarization sensitivity, 19, 208, 214, 222, 248, 251, 252 Polychaete, 31 Polychelida, 3, 13 Polyphemus, 208, 434 Pontella, 211 Porcellio, 23 Portunus, 118 Postexcitatory inhibition, 402, 404 Postinhibitory rebound, 114, 122, 125, 131, 379 Postsynaptic inhibition, 96, 102, 105, 151, 152 Posture, 6, 7, 20, 68–70, 74, 86, 87, 117, 225, 294, 362, 363, 366, 367, 372, 373, 377, 379, 381, 384, 386, 388, 412, 414, 415, 422, 459, 460, 462, 466, 469, 548 Potamobius, 436

562 Index Predation, 8, 17–20, 31, 32, 49, 50, 59, 63, 64, 66, 69, 75, 209, 210, 235, 236, 239–242, 244, 245, 248, 249, 252, 255, 263–265, 285, 293, 294, 296, 297–303, 313, 314, 396, 399–401, 410–413, 418, 421, 484, 485, 487, 490–492, 494, 497–502, 512, 513, 517 Predator avoidance, 255, 484, 485, 487, 499–502 Presynaptic inhibition, 86, 93, 96, 97, 105, 116, 123, 124, 147, 148, 151–153, 166, 277, 382, 384, 386–388, 416 Prey, 12, 17, 18, 29, 49, 59, 66, 69, 235, 241, 242, 244, 248, 249, 265, 272, 280, 281, 283, 293, 294, 296, 298, 300, 302, 303, 314, 331, 387, 396, 412, 492, 512, 514 Procambarus, 19, 20, 22, 25, 26, 28, 33, 51, 52, 56, 62, 68, 71, 74, 94, 117, 178, 183, 189, 192, 208, 211, 213, 226, 251, 363, 365, 431–434, 436, 438, 439, 444, 461, 465, 468 Proctolin, 106, 107, 120, 121, 123, 125–130, 345, 376 Projection neuron tract, 37 Proliferation zone, 175, 182–188, 193 Proprioception, 73, 378, 381, 549 Proprioceptor, 18, 49, 66, 67, 69, 70, 72, 74, 152, 225, 367–371, 373, 376, 381, 384, 386, 543 Propus, 364, 365, 371 Protein synthesis, 158, 163, 165, 346, 519, 520 Protocerebral tract, 16, 37, 184, 185, 268, 437, 438, 446, 447, 462, 521, 521, 528 Protocerebrum, 12, 14–17, 20, 21, 33, 35–40, 54, 57, 58, 60, 185, 186, 206, 218, 219, 221, 223–227, 268, 278, 279, 438, 441, 444–447, 522–524 lateral, 14, 16, 17, 21, 33, 35, 37, 54, 57, 185, 186, 218, 219, 221, 223–225, 268, 278, 279, 444, 446, 447, 523, 524 medial, 16, 224 median, 14, 20, 33, 185, 444, 445, 447 Proximal glial sheath, 446 Pseudopupil, 210, 237–240, 441, 493, 495, 496 Pterygota, 37, 39 Pugettia, 267 Pullosquilla, 243, 254 Pycnogonida, 3 Pyloric region, 122, 338, 343 Receptor anterior gastric (AGR), 123, 132, 133, 341, 342, 344 bend, 304, 309 bimodal, 29, 308 gastropyloric (GPR), 116, 123, 124, 127, 131, 133, 344 posterior stomach, 344 scolopidial, 305

stretch, 66, 74, 117, 149, 151, 294, 331, 344, 368, 401, 543, 544, 547 Receptor muscle, 72, 368, 369 Receptor potential, 70, 73, 212, 216, 217, 226, 227, 274, 439, 441 Reflex, 2, 12, 18, 20, 21, 33, 52, 54, 63, 66, 69, 70, 73, 74, 148, 206, 212, 223–226, 228, 249, 372, 377, 381–388, 397, 400, 405, 406, 408, 410, 412, 413, 416, 417, 437, 511, 540 Reflex reversal, 381–384, 386, 388 Reflex, oculomotor, 21, 224 Refractive index, 59, 242, 243 Remipedia, 236 Reptantia, 3, 13, 14, 26, 30, 37–39, 41, 70, 74, 177, 186, 278, 299, 313 Resistance reflex, 73, 74, 373–377, 381, 382, 384, 385, 387, 388 Resting potential, 100, 132, 342, 387, 537, 540, 547, 550 Retina, 15–21, 64, 210, 212, 213, 218, 228, 236, 237, 241–244, 247, 252, 254, 255, 257, 428, 434, 436–439, 441, 444–448, 493, 521 Rhabdomere, 59–63, 207, 213, 215, 247, 249 Rhithropanopeus, 238 Rhythm biological, 6, 7, 429, 434 cardiac sac, 128, 129, 338 circadian, 58, 191, 195, 212, 227, 249, 256, 428–445, 447–449 diurnal, 25, 32, 49, 207, 209, 210, 429, 430, 434, 435, 440, 548 gastric mill, 123, 124, 129, 136, 337–339, 343, 345, 349–350 infradian, 429, 430 pyloric, 122, 123, 127, 130–133, 135, 136, 337–339, 343–346, 348, 349–350 ultradian, 429, 434 Rhythmic bursting, 114, 116, 120, 127, 128, 138, 276, 342 Rimicaris, 212, 242 RNA interference, 5, 8 Sarcomere, 86, 91, 97–100, 102, 104, 539 Sarcoplasmic reticulum, 97, 102, 120 Scaphognathite, 22, 86 Scolopale, 51, 52, 54, 74, 178, 266, 270, 271, 305–310, 314 Scylla, 70, 72, 221 Scyllaridae, 22, 177 Sensilla, beaked, 51, 52, 54 Sensilla, hooded, 22, 51–53, 268–270 Sensitization, 150, 416, 511, 513 Sensory cone, 24

Index 563 Sensory fiber, 11, 368, 369 Serotonin, 106, 107, 116, 126–128, 131, 135, 189, 190, 195, 344, 346, 374, 376, 377, 388, 412, 412, 438–440, 458, 465, 467–471, 511, 545, 546 Sesarma, 25, 26, 435, 442 Setules, 51, 269, 304, 313 Shelter, 5, 23, 31, 32, 235, 264, 270, 285, 321, 458, 459, 465, 467–469 Shrimp brine, 3, 303 caridean, 3, 69, 242, 299, 469, 474, 475 cleaner, 27, 29, 30 fairy, 3, 303 freshwater, 463, 467, 472 mantis, 3, 13, 17, 20, 63, 75, 240, 245, 247, 250, 252, 418, 466, 467 sergestid, 54, 304, 313 snapping, 75, 467 Sicyonia, 187 Signal transmission, 535, 549 Sinus gland, 116, 122, 224, 345, 437, 441, 448 Skogsbergia, 212 Social interaction, 5, 12, 31, 33, 49, 63, 281, 363, 485, 490, 502 Social status, 6, 7, 22, 195, 264, 363, 388, 414, 415, 421, 422, 457, 458, 460, 462, 466–468, 470, 472, 473, 475 Somata, 2, 15, 33, 34, 35, 55, 74, 120, 156, 175, 182–187, 191, 198, 221, 277, 305, 311, 338, 339, 341, 342, 345, 347, 366, 444, 448, 474, 522, 524 Spectral sensitivity, 58, 206, 207, 213, 214, 228, 236, 243, 247, 248, 255, 256, 494 Spermatophore, 22 Spike frequency regulation, 114 Sponge, 31 Squilla, 254, 420 Squillidae, 18 Statocyst, 12, 69, 70, 72, 75, 76, 181, 225, 226, 301, 363, 548 Statocyst cavity, 70, 363 Stenopodidea, 3, 13, 14, 29, 177, 278 Stenopus, 15, 27, 29 Stimulus chemical, 21, 25, 52, 265, 271, 273, 276, 277, 280, 281, 283 mechanical, 21, 52, 224, 299, 308, 309, 313, 367, 402, 406, 435, 525 visual, 12, 20, 33, 35, 226, 461, 518, 525, 545 Stomatogastric ganglion, 6, 86, 114, 115, 122, 134, 149–151, 154, 158, 337–339, 342, 347, 366 Stomatogastric nervous system, 122, 151, 337, 338, 340, 350

Stomatopoda, 2, 3, 11, 13–15, 18, 19–21, 27–31, 40, 49, 59, 61–65, 76, 207–210, 214, 215, 218, 221, 222, 228, 235, 238–240, 242, 243, 245–248, 250–255, 257, 266, 278, 418–420, 465, 512 Stomodeum, 16 Superposition, 59, 61, 62, 207–213, 237, 238, 240, 241, 253, 254, 256 Supraesophageal ganglion, 185, 212, 224, 411, 436, 441, 442, 521 Swimmeret, 75, 114, 115, 117, 118, 121, 151, 366, 368, 373, 399, 417, 419, 541, 549 Synaptic transmission, 86, 108, 115–117, 135, 148, 166, 342, 376, 396, 401, 407, 440, 535, 537, 538, 540, 550 Syncarida, 3, 14, 18, 418 Synchelidium, 435 System, sensory, 5, 6, 8, 12, 30, 33, 49, 50, 75, 76, 248, 255, 274, 285, 301, 313, 362 System, visual, 17–21, 25, 27, 38, 40, 63, 64, 76, 206, 221, 225, 235, 236, 239, 243, 244, 248–250, 253, 255, 257, 332, 485, 492, 494, 495, 513, 520 Tail flip, 7, 73, 149, 158, 229, 366, 396–400, 406–412, 414, 416–418, 420, 421, 423, 460, 462, 464, 466, 510 Talitrus, 323, 324, 433, 435 Talorchestia, 238, 249, 324 Tapetum, 59, 60, 207, 210–212 Telson, 12, 63, 67–69, 71, 72, 178, 227, 250, 401, 407, 417, 420, 541 Temora, 309 Terminal medulla, 11, 12, 14–16, 21, 33–38, 41, 42, 57, 184, 185, 278, 279, 442, 444–448, 474 Territorial behavior, 499 Tetraconata, 176, 177, 179–181, 190, 196 Thalamita, 512 Thalassinida, 26, 38 Thoracic ganglion, 54, 73, 331, 369, 379, 380, 384, 443 Thorax, 397, 405, 411, 416, 459, 545 Thorax-coxa muscle receptor organ (TCMRO) 368–370, 375, 381–383 Threshold, 41, 50, 59, 63, 66, 68, 75, 100–103, 105, 109, 116, 128, 131, 132, 215, 216, 273, 275, 280, 283, 296, 300, 301, 311–313, 342, 366, 367, 374, 382, 384, 403–405, 408, 412, 413, 417, 419, 431, 466, 470, 501, 538, 543, 547 Thysanoessa, 253 Thysanopoda, 253 Tigriopus, 264 Transcuticular sensillum, 67 Transmission, chemical, 1, 342

564 Index Tritocerebrum, 12, 14, 15, 16, 17, 29, 37, 54, 69, 185, 446, 447 Turbidity, 17, 25, 303, 325 Turbulence, 270, 280–283, 297, 301, 314, 416 Uca, 25, 26, 63, 207–209, 237–239, 248–250, 267, 272, 273, 433–435, 440, 486–491, 494, 497, 499, 500, 513 Undinula, 300 Urocaridella, 29 Uropod, 63, 67, 69, 73, 86, 227, 368, 373, 401, 411, 420 Vision color, 59, 63, 206, 213, 214, 227, 248, 249, 251, 252, 494 contrast, 206 polarization, 206, 210, 221, 239, 247, 248, 251, 252 Visual danger stimulus (VDS), 509, 513–517, 519, 520, 524, 525, 527

Visual pigment, 63, 213, 243–249, 251, 253–257, 494 Volcanic vent, 13 Walking, 2, 4–7, 12, 19, 27, 31, 32, 41, 52, 58, 73, 86, 88, 90, 93, 97, 99, 105, 106, 115, 118, 149, 159–163, 181, 227, 268, 294, 309, 325, 327, 328, 330, 331, 362–366, 368, 371–373, 375, 377–381, 384–388, 411, 459, 460, 462, 463, 487, 489, 490, 497, 499, 502, 503, 512, 538, 544, 547 Water deformation, 296, 304, 313, 314 Wavelength, 7, 40, 59, 63, 214, 215, 218, 243–250, 254, 255, 264, 494 Waving display, 498, 503 Xenocarida, 3 X-organ, 116, 122, 221, 224, 437, 439, 440, 448 Zooplankton, 210, 255, 294