Developmental Biology and Larval Ecology: The Natural History of the Crustacea, Volume 7 [7] 0190648953, 9780190648954

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Developmental Biology and Larval Ecology

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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 Physiology (Volume 4) Edited by Ernest S. Chang and Martin Thiel Life Histories (Volume 5) Edited by Gary Wellborn and Martin Thiel Reproductive Biology (Volume 6) Edited by Rickey Cothran and Martin Thiel Developmental Biology and Larval Ecology (Volume 7) Edited by Klaus Anger, Steffen Harzsch, and Martin Thiel

Developmental Biology and Larval Ecology The Natural History of the Crustacea Volume 7

EDITED BY KLAUS ANGER, STEFFEN HARZSCH, 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 is a registered trade mark 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, United States of America. © Oxford University Press 2020 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 Control Number: 2020907951 ISBN 978–0–19–064895–4 9 8 7 6 5 4 3 2 1 Printed by Integrated Books International, United States of America

PREFACE

This is the seventh volume of a 10-​volume series on The Natural History of the Crustacea. This volume examines developmental biology and larval ecology, and it follows Volume 1: Functional Morphology and Diversity, Volume 2:  Life Styles and Feeding Biology, Volume 3: Nervous Systems and Control of Behavior, Volume 4: Physiology, Volume 5: Life Histories, and Volume 6: Reproductive Biology. The remaining three volumes will explore additional aspects of crustacean natural history, evolution and biogeography, fisheries and aquaculture, and ecology and conservation biology. Chapters in this volume synthesize our current understanding of early crustacean development from the egg through the embryonic and larval phases. The first part of this volume focuses on the elemental aspects of crustacean embryonic development. Gerhard Scholtz describes early embryonic processes such as cleavage and gastrulation, which form the germ band that subsequently structures into segmental units and grows by the addition of new segments. The next chapter is devoted to limb morphogenesis; Anastasios Pavlopoulos and Carsten Wolff explain how crustaceans unfold their body plan and how the appendages develop, including aspects of regeneration. The development of the internal organs in crustaceans is a next major step in their ontogeny, and the chapter by Günther Loose and his collaborators is devoted to highlighting these processes. Developmental malformations are frequently observed in crustaceans, and Gerhard Scholtz describes the range of deformations known in crustaceans and discusses possible factors that may cause such malformations. Finally, in the last chapter of this section, Martin Fritsch and colleagues explain the processes that lead to hatching and free-​living individuals. The second part of the book provides an account of the larval phase of crustaceans and describes processes that influence the development from hatching to an adult-​like juvenile. Patterns of postembryonic development following hatching are described by Ole Sten Møller and his collaborators, and the impacts of environmental conditions on the development of the larvae are explored by Chaoshu Zeng and colleagues. At the end of their planktonic phase, benthic crustaceans face multiple challenges to find suitable habitats, which involve diverse settlement strategies; these are examined by Paulina Gebauer together with her collaborators. During settlement, these larvae experience variable degrees of structural transformations, generating some of the most diverse morphologies observed in nature; Joachim Haug explores the variable degrees of metamorphosis in the main crustacean clades. Luis Giménez highlights how phenotypic plasticity and, in particular, maternal traits affect larval development in crustaceans; how larvae with different developmental histories respond to prevailing environmental conditions; and how the larval history can affect later life history phases (carryover effects).

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vi Preface The third and final part of the book explores ecological interactions during the planktonic phase and how crustacean larvae manage to find food, navigate the dynamic water column, and avoid predators in a medium that offers few refuges. Aspects of nutrition and feeding of planktonic larvae are discussed by Andrew Jeffs and Richard O’Rorke. Jonathan Cohen and Charles Epifanio describe how crustacean larvae react to a wide variety of environmental stimuli from the liquid medium that carries them throughout their planktonic life. Being continuously exposed to predators and competitors is a major challenge for the growing larvae, leading to astonishing morphological and behavioral adaptations that are reviewed by Samuel Bashevkin and Steven Morgan; Steven also examines the processes that influence dispersal of the planktonic larvae. Applying current knowledge of larval life and dispersal, Per-​Olav Moksnes and Per Jonsson explore how this information can be used to investigate the connectivity of crustacean populations and design effective marine protected area networks. Together with the chapters on metamorphosis, settlement, and carryover effects of variation in larval condition, this final chapter reconnects the planktonic larvae to the benthic adults, thereby completing the cycle and the volume. Collectively, these 15 chapters provide a thorough overview of our current knowledge across the major themes in crustacean developmental biology, including larval ecology. We expect this volume to be valuable to scholars and students who are interested in gaining deeper insights into the processes that lead from a single cell to subsequent stages of life and how the growing organisms face the challenges posed by their environment. We hope this synthesis and the thoughtful ideas provided by the expert contributors to this volume spur new avenues of research on developmental biology and larval ecology within the Crustacea and beyond.

ACKNOWLEDGMENTS

We thank our contributors for graciously sharing their time, energy, knowledge, and insights to make this volume possible. It has been both a pleasure and honor to work with each of them. Our editorial assistants, Annie Mejaes, Mika Tan, Tim Kiessling, Miles Abadilla, and Miguel Angel Penna-Díaz, were impeccably skilled and organized, and always kept us moving forward. We thank our external reviewers for their valuable and generous feedback. Foremost, we are grateful to all our contributors for their time and patience; they have made an extra effort in this fast-​paced time of scientific study to provide excellent overviews that will prove useful to active and incoming scholars of modern crustacean natural history. Finally, we express our appreciation to our publisher, Oxford University Press, for their commitment to this project. Editing of this book was generously supported by Universidad Católica del Norte, Chile.

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CONTRIBUTORS

EDITORS Klaus Anger Malente, Germany Steffen Harzsch Zoological Institute and  Museum University of Greifswald Greifswald, Germany Martin Thiel Facultad Ciencias del Mar Universidad Católica del Norte Coquimbo, Chile AUTHORS Samuel M. Bashevkin Bodega Marine Laboratory University of California Davis Bodega Bay, CA, United States of America Guy Charmantier University of Montpellier Montpellier, France Mireille Charmantier-​Daures University of Montpellier Montpellier, France

Jonathan H. Cohen School of Marine Science and Policy University of Delaware Lewes, DE, United States of America Charles E. Epifanio Harrington Professor Emeritus School of Marine Science and Policy University of Delaware Lewes, DE, United States of America Martin Fritsch Museum für Naturkunde Berlin Leibniz Institute for Research on Evolution and Biodiversity Berlin, Germany Paulina Gebauer Universidad de Los Lagos Puerto Montt, Chile Luis Giménez School of Ocean Sciences Bangor University Anglesey, United Kingdom Alfred-Wegener-Institut , Helmholtz Center for Polar and Marine Research, Helgoland Marine Station Helgoland, Germany

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List of Contributors Guillermo Guerao Departament de Ciències Institut Banús (Departament d’Educació) Barcelona, Spain

Per-​Olav Moksnes Department of Marine Sciences University of Gothenburg Gothenburg, Sweden

Joachim T. Haug Department of Biology II Zoomorphology University of Munich (LMU) Planegg-Martinsried, Germany

Ole Sten Møller Campus Bornholm STX Section Rønne, Denmark

Iván A. Hinojosa Departamento de Ecología Facultad de Ciencias Universidad Católica de la Santísima Concepción Concepción, Chile Centro de Investigación en Biodiversidad y Ambientes Sustentables (CIBAS) Universidad Católica de la Santísima Concepción Concepción, Chile Millennium Nucleus for Ecology and Sustainable Management of Oceanic Islands (ESMOI) Departamento de Biología Marina Facultad de Ciencias del Mar Universidad Católica del Norte Coquimbo, Chile

Steven G. Morgan Bodega Marine Laboratory University of California Davis Bodega Bay, CA, United States of America Richard O’Rorke Leigh Marine Laboratory Institute of Marine Science University of Auckland Warkworth, New Zealand Jørgen Olesen Natural History Museum of Denmark University of Copenhagen Copenhagen, Denmark

Andrew Jeffs Institute of Marine Science School of Biological Sciences University of Auckland Auckland, New Zealand

Kurt Paschke Instituto de Acuicultura Universidad Austral de Chile Puerto Montt, Chile Centro FONDAP de Investigación en Dinámica de Ecosistemas Marinos de Altas Latitudes (IDEAL) Valdivia, Chile

Per R. Jonsson Department of Marine Sciences—​Tjärnö University of Gothenburg Strömstad, Sweden

Anastasios Pavlopoulos Institute of  Molecular Biology and Biotechnology Foundation for Research and Technology Hellas Greece

Günther J. Loose Department of Biology, Comparative Zoology Humboldt University of Berlin Berlin, Germany

Nicholas Romano Department of Aquaculture and Fisheries University of Arkansas at Pine Bluff Pine Bluff, AR, United States of America

Guiomar Rotllant Departament de Recursos Marins Renovables Institut de Ciències del Mar—​CSIC Barcelona, Spain Spain Gerhard Scholtz Department of Biology, Comparative Zoology Humboldt University of Berlin Berlin, Germany Günter Vogt Faculty of Biosciences University of Heidelberg Heidelberg, Germany

List of Contributors Carsten Wolff Department of Biology, Comparative Zoology Humboldt University of Berlin Berlin, Germany Chaoshu Zeng College of Science & Engineering James Cook University Townsville, Australia

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CONTENTS



1. From a Single Cell to Segmental Structures: Crustacean Embryology  •  1 Gerhard Scholtz



2. Crustacean Limb Morphogenesis during Normal Development and Regeneration  •  45 Anastasios Pavlopoulos and Carsten Wolff



3. Organogenesis • 79 Günther Loose, Günter Vogt, Mireille Charmantier-​Daures, Guy Charmantier, and Steffen Harzsch



4. Duplicated, Twisted, and in the Wrong Place: Patterns of Malformation in Crustaceans • 112 Gerhard Scholtz



5. Hatching • 143 Martin Fritsch, Jørgen Olesen, Ole Sten Møller, and Günther Loose



6. Patterns of Larval Development  •  165 Ole Sten Møller, Klaus Anger, and Guillermo Guerao



7. Effects of Environmental Conditions on Larval Growth and Development  •  195 Chaoshu Zeng, Guiomar Rotllant, Luis Giménez, and Nicholas Romano



8. Settlement and Metamorphosis in Barnacles and Decapods  •  223 Paulina Gebauer, Luis Giménez, Iván Hinojosa, and Kurt Paschke



9. Metamorphosis in Crustaceans  •  254 Joachim T. Haug



10. Phenotypic Plasticity and Phenotypic Links in Larval Development  •  284 Luis Giménez

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11. Feeding and Nutrition of Crustacean Larvae  •  309 Andrew Jeffs and Richard O’Rorke



12. Response to Visual, Chemical, and Tactile Stimuli  •  332 Jonathan H. Cohen and Charles E. Epifanio



13. Predation and Competition  •  360 Samuel M. Bashevkin and Steven G. Morgan



14. Dispersal • 383 Steven G. Morgan



15. Larval Connectivity and Marine Protected Area Networks  •  408 Per-​Olav Moksnes and Per R. Jonsson Index • 437

Fig 1.2

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Total and mixed cleavage of various crustaceans with a stereotyped cell division pattern. (A) The amphipod Cryptorchestia garbinii. (B) The euphausiid Meganyctiphanes norvegica. (C) The dendrobranchiate decapod Sicyonia ingentis. (D) Dorsal aspect of the cirriped Austrominius modestus. (Dʹ) Ventral aspect. (E) Vegetal aspect the cladoceran Bythotrephes longimanus. (Eʹ) Lateral aspect. Sister cells are connected with a black line. Except for the amphipod C. garbinii yellow marks the A quadrant; red, the B quadrant; green, the C quadrant; and blue, the D quadrant, indicating putative homologies. The nomenclature of individual cells is only added in B. longimanus (en, cells forming the endoderm, g, the primordial germ cells). The cleavage of Amphipoda is so different from the other crustaceans that the coloring is chosen arbitrarily. The arrows point to the gastrulation center. In malacostracans, it occurs at the end of a cell band (blue blastomeres meeting the center of the yellow–​green band); in non-​malacostracans, in the center of a cell band (contact zone of the blue and red blastomeres). Each column shows, from top to bottom, the four-​cell stage, the eight-​cell stage, the 16-​cell stage, and the 32 cell-​stage (the latter is not, in all cases, complete). B. longimanus undergoes a mixed cleavage in which the early stages are characterized by a central yolk mass. C. garbinii, A. modestus, and B. longimanus display unequal cleavages (micromeres and macromeres), whereas those of M. norvegica and S. ingentis are equal. The two interlocking cell bands (tennis ball pattern) is best seen in the euphausiacean and the decapod. In the cirriped, it is slightly transformed in the B and D quadrants; in the cladoceran, the contact zone (cross-​furrow) of the B and D quadrants is found at the vegetal pole, and of the A and C quadrants at the animal pole, as in the cirriped, despite an arrangement lacking the characteristic cell bands. (A) Modified after Scholtz and Wolff (2013, Fig. 4.4) and Scholtz and Wolff (2002, Fig. 4c). (B) Modified after Alwes and Scholtz (2004, Fig. 5). (C) Modified after Hertzler and Clark (1992, Fig. 6). (D, Dʹ) Modified after Ponomarenko (2014, Fig. 3.3.1). (E, Eʹ) Modified after Alwes and Scholtz (2014, Fig. 7).

   

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Fig. 1.3.

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Superficial cleavage. (A–​F) From the two nucleus stage to the 32-​cell stage of the cleavage of the isopod Idotea. The spindle axes leading to the four-​nuclei stage show a 90-​degree angle to each other. The nuclei reach the egg’s surface between the 16-​and the 32-​cell stages. Cell boundaries are formed at the 32-​cell stage. (G) Four-​nuclei stage of the isopod Porcellio scaber with the same arrangement of the nuclei (dark areas) within the yolk mass as in (B). (H–​K) Indications of cell fate determination in superficial cleavage. (H) Transition between the 32-​and 64-​cell stages of the cumacean Diastylis rathkei. Two cells show a retarded division and are the first to form cell boundaries. These probably mark the future gastrulation center. (I–​K) Sixteen-​to 64-​cell stages of the cladoceran Leptodora kindtii. In this case, superficial cleavage forms the pattern of two interlocking bands comparable with that of some total cleavages (see Fig. 1.2). (A–​F) Modified after Strömberg (1965, Fig. 1). (G) Modified after Scholtz and Wolff (2013, Fig. 4.5). (H) Modified after Dohle (1970, Fig. 5). (I–​K) Modified after Alwes and Scholtz (2014, Fig. 10).

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Fig. 1.5.

Cell lineage studies. (A, B) The amphipod Cryptorchestia garbinii. (C, D) The cirriped Austrominius modestus. (E) The decapod Sicyonia ingentis. (A) Live images of the cleavage from the zygote to early gastrulation. The image of the 16-​ cell stage in the center is a scanning electron microscope micrograph. (B) The result of the single-​cell application of the fluorescent dye DiI to two micromeres (shown in red in the inset). The entire postnaupliar mesoderm, including the mesoteloblasts plus the endodermal midgut glands, show the staining. (C) Ventral (left) and dorsal (right) aspects of the transition of the 16-​to 32-​cell stage (Sytox green marker of cell nuclei). (D) Maps of the contributions of the quadrants to the external ventral (left) and dorsal (right) sides and internal organs (nervous system, musculature, intestine) of the nauplius larva. (E) Cell lineage markings of the blastomeres of the four-​cell stage and their outcome (dotted areas) in the nauplius larva. The upper row in each box shows the ventral side; the lower row, the dorsal side. Mirror images are the results of the two chiral (mirror image) variants of eggs. (A) Modified after Scholtz and Wolff (2002, Fig. 1). (B) Modified after Wolff and Scholtz (2002, Fig. 12). (C, D) Modified after Ponomarenko (2014, Figs. 3.3.6 and 3.4.1). (E) Modified after Hertzler and Clark (1992, Fig. 6) and Hertzler et al. (1994, Fig. 4).

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Fig. 1.7.

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Gastrulation. Modes of endoderm, mesoderm, and germ cell separation. (A–​F) Superficial aspects of the gastrulation center. (G–​K) Sections through the gastrulation center. (A) The anterior end of the germ band is characterized by the head lobes (hl), posterior invagination with a blastopore (bp) close to the proctodaeum (pr) and a large endoderm (en) sack [compare with (K)]: crayfish Cherax destructor. (B) Epiboly with micromeres overgrowing (dotted line) the yolky macromere: cirriped Austrominius modestus. (C) Immigration with a slight invagination and early determination of the primordial germ cells (the arrow points to the germ granules) (green), the endoderm (yellow), and the crown cells forming mesoderm (red): cladoceran Polyphemus pediculus. (D) Anterior invagination forming a blastopore (bp) combined with immigration [compare with ( J)]: amphipod Cryptorchestia garbinii. (E) Late gastrulation with immigration and formation of ectodermal teloblasts (ET) arranged in a semicircle in front of the gastrulation center (ga): isopod Asellus communis. (F) Immigration of two large mesendoderm cells surrounding crown cells (cc) (mesoderm) with radial mitotic spindles, and subsequent invagination forming a blastopore (bp):  euphausiacean Meganyctiphanes norvegica. (G) Immigration of two large mesendoderm cells (me) [one containing the intracellular body (icm), a kind of germ granule] surrounding crown cells and, later, invagination:  dendrobranchiate Penaeus monodon. (H)  Three subsequent stages (from left to right) (sagittal section). Gastrulation center (ga) with immigration forming a shallow pit. The ectodermal teloblasts (ET) differentiate at the anterior margin of the gastrulation center and produce smaller ectoderm cells in an anterior direction, primordial germ cells (gc), mesoderm (mes), and yolk containing endoderm cells (en) migrate anteriorly, mesoteloblasts (MT) bud further mesoderm cells in the anterior direction [compare with (Figs. 1.9 and 1.10)]: mysid Hemimysis lamornae. (I) Sagittal section of later stage of (C) with an almost closed blastopore (bp), showing immigration of germ cells (gc), mesoderm (mes), and endoderm (en). ( J) Cross-​section of (D) depicting invagination (bp) and cell immigration (asterisks). (K) Sagittal section of Astacus astacus; compare with (A). The endoderm (a) forms a large midgut primordium with secondary yolk pyramids, the proctodaeum (pr) elongates and forms the hindgut, and the invagination of the stomodaeum (st) begins. The large ectoteloblasts (ET) surrounding the caudal papilla and some mesoderm cells are visible. (A) Photo courtesy of Gerhard Scholtz. (B) Photo courtesy of Ekaterina Ponomarenko. (C, I) Modified after Kühn (1913, Figs. 50 and 74). (D) Photo courtesy of Gerhard Scholtz. (E) McMurrich (1895, Fig. 36). (F) Photo courtesy of Frederike Alwes. (G) Modified after Biffis et al. (2009, Fig. 7f). (H) Modified after Manton (1928, Fig. 3). ( J) Modified after Scholtz and Wolff (2002, Fig. 12b). (K) Modified after Reichenbach (1896, Fig. 66).

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Fig. 1.8.

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Egg–​nauplius and nauplius. (A) The egg–​nauplius of the crayfish Astacus astacus with the advanced anlagen of the eyes (head lobes, hl), the first and second antennae and the mandibles (a1, a2, md) compared with more posterior segmental structures. At the anterior, there is the anlage of the labrum (lr); at the posterior, there is the forming caudal papilla (cp) with the proctodaeum. (B) The nauplius larva of the northern krill Meganyctiphanes norvegica. This is a reduced nonfeeding nauplius with few cells (nuclear fluorescent stain bisbenzimide). (C) Scanning electron micrograph of an advanced embryo of the amphipod Gammarus pulex. Here an egg–​nauplius is lacking because the naupliar region shows no advanced development. The naupliar appendages are in a similar stage as the crayfish in (A); but, in contrast to this, the fifth thoracic segment shows early limb buds and the fourth pleonic segment is recognizable. cf, caudal furrow. (D) Scanning electron micrograph of the nauplius II of the cirriped Amphibalanus improvisus showing the advanced development of a proper nauplius compared with an egg–​nauplius (B) and the nauplius of the krill (B). A large labrum (lr) is present. ff, frontal filaments. (A) Modified after Reichenbach (1896, Fig. 9). (B) Photo courtesy of Gerhard Scholtz. (C) Photo courtesy of Gerhard Scholtz. (D) Modified after Semmler et al. (2009, Fig. 1b).

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ie

Fig. 1.9.

(B) Germ band of Neomysis integer. There are relatively few cells, which are arranged regularly in a grid-​like pattern of longitudinal columns (including a midline) and transverse rows. The large ectoteloblasts (arrow) give rise to regular rows that follow a determined sequence of division (yellow: undivided rows, blue: rows in the phase of the first wave of division, green: rows in the phase of the second wave of division, orange: beginning differential cleavages and morphological segmentation, red: the offspring of one ectoteloblast along the anteroposterior axis of the germ band). ie, intercalary elongation; sb, segmental border; gb, genealogical boundary (for an explanation, see the text).

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Fig. 1.10.

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Teloblasts. (A) A cross-​section through the caudal papilla in the area of teloblasts of Homarus americanus. An outer circle of 19 ectoteloblasts (ET) and an inner circle of eight mesoteloblasts (MT) around the proctodaeum (pr) are visible. This is the ancestral condition within Malacostraca. (B) A similar section in the freshwater crayfish Cherax destructor. About 40 ectoteloblasts are combined with eight mesoteloblasts, both situated around the proctodaeum (pr). (C) The growth zone of the peracarid Neomysis integer. The ectoteloblasts form a transverse row of about 15 cells. (D) The germ band with ventrally folded caudal papilla of Astacus astacus. The arrowpoints to the ectoteloblasts. Again, these are more than 19 in number and Reichenbach (1886) is the first record of this type of stem cell. (E) Ventral aspect of the germ band of the isopod Porcellio scaber. In this case, the germ band growth around the yolk occurs without the formation of a caudal furrow or a caudal papilla. (F) The cell division pattern of the differentiation of ectoteloblasts in some decapod crustaceans. (G) The cell division pattern of the differentiation of mesoteloblasts in the peracarid Diastylis rathkei. A corresponding pattern has been described for amphipods and decapods. (A) Modified after Dohle et al. (2004, Fig. 3). (B) Photo courtesy of Gerhard Scholtz. (C) Photo courtesy of Gerhard Scholtz. (D) Modified after Reichenbach (1896, Fig. 10). (E) Modified after Scholtz and Wolff (2013, Fig. 4.6c). (F) Modified after Ooishi (1959, Fig. 8). (G) Modified after Dohle et al. (2004, Fig. 5a).

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Fig. 1.11.

Segment formation and segment polarity engrailed expression in Cryptorchestia garbinii. (A) Differential interference contrast image of the whole mount of a germ band showing a thoracic segment during formation. The expression of engrailed is made visible with an antibody (brown nuclei). Each transverse stripe including the midline (m) marks the posterior margin of a segment. The intersegmental furrows form posterior to the engrailed expression. The stereotyped arrangement of ectoderm cells is recognizable. (B) Drawing of the preparation of (A) with an analysis of the mitoses of the differential cleavages and the clonal composition. Lines connect sister cells. (C) Scanning electron micrograph of three segments of the embryonic thorax (th) with artificial blue staining of the cells that express engrailed. This technique demonstrates the morphogenesis of the segmental furrows and the limb buds. Lines connect sister cells. The labels mark individually identified cells. ml, midline. (A–​C) Modified after Dohle et al. (2004, Figs. 6a and 12).

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Fig. 1.12.

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The cumacean Diastylis rathkei. Again, this morphogenesis correlates with the differential cleavages of the malacostracan germ band. The arrow marks the genealogical boundary between the descendants of two ectoderm rows. (B) Scheme of the expression of distal-​ less in a thoracic uniramous and a pleonic biramous limb of Cryptorchestia garbinii. The empty circles indicate the loss of a previous expression. This exemplifies the similar anlage of uniramous and biramous limb buds and the later differentiation resulting from a loss of distal-​ less expression. (C) Clonal analysis of uniramous thoracic and biramous pleonic limbs of the same species using the fluorescent vital marker DiI. All clones found in in the biramous limb along the long axis from the protopod (prp) to the tips of the exo-​and endopods (exo, endo) are distributed similarly in the uniramous limb (cx, coxa; ba, basis; is, ischium; me, merus; ca, carpus; pro, propodus; da, dactylus), suggesting that uniramous limbs are the result of the suppression of the bifurcation of the limb buds. The exo-​ and endopods of limbs are formed by the same proximodistal axis. Exites such as gills (gi) and coxal plates (cxp) are the result of additional axis formations. The cells adjacent to the midline (light and dark gray 1,2) form part of the sternite (ster); those at the laterodorsal margin of the germ band (blue and lilac 8,9) contribute to the tergite (ter). (D) Embryo of the branchiopod Cyclestheria hislopi with distal-​ less expression in the lobes of the limb anlagen, the labrum, and the furca. (A, B) Modified after Dohle et al. (2004, Figs. 10 and 14). (C) Modified after Wolff and Scholtz (2008, Figs. 3e and 4e). (D) Photo courtesy of Gerhard Scholtz.

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Fig. 1.13.

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(E)

(G)

Early neurogenesis. (A) Sagittal section through the nauplius of the copepod Tigriopus californicus. Anterior to the left. The forming segmental ganglia (dotted lines) are depicted and the first longitudinal fiber tracts (connectives) (arrowhead) are visible. At the ventral side, neuroblasts (asterisks) are visible. Neural stem cells that produce offspring (ganglion mother cells) are arranged as columns into the interior of the larva. In the end, the nerve cells (dots) differentiate. (B) Ventral aspect of the anlage of a segment of the amphipod Gammarus pulex during differential cleavages that lead to the differentiation of superficial neuroblasts, which in turn give rise to ganglion mother cells. In the left body, half the superficial cells are omitted and only ganglion mother cells are visible. The median neuroblast is also omitted. (C–​E) Neuroblasts in the amphipod Cryptorchestia garbinii. (C) The neuroblast map (ventral view) with all identified lateral neuroblasts. The median neuroblast is omitted. (D, E) The clone of an identified neuroblast (b1hn) that generates the pioneer neurons pCC and aCC, known from Drosophila. (D) Schematic drawing. (E) The preparation with DiI. ac, anterior commissure; pc, posterior of a ganglion. Other labels indicate identified individual cells after differential cleavages as in Fig. 1.11. (F, G) Neuroblasts in the branchiopod Triops cancriformis. (F) Transverse section through a larval trunk segment. Ventral side at the bottom, with lateral neuroblast (arrows) and a median neuroblast. On top of each neuroblast there are columns of smaller ganglion mother cells. (G) Scheme of neuroblasts (asterisks) and the columns of smaller ganglion mother cells. Gray: mesoderm. (A) Modified after Hein and Scholtz (2018, Fig. 2b). (B) Modified after Dohle et al. (2004, Fig. 9). (C–​E) Modified after Ungerer and Scholtz (2008, Figs. 3 and 5). (F, G) Modified after Müller (2006, Figs. 12a and 15).

(A)

(B)

Fig. 4.2.

Experimentally produced heteromorphoses by artificially misexpression of the ubx gene in the amphipod Parhyale hawaiensis. (A)  A  thoracopod replacing the right first antenna (highlighted in blue). (B)  Transformation of the segments of the second maxilla and the maxilliped to anterior pereopod segments including coxal plates and the transformation of anterior walking legs to a more posterior morphology (highlighted in blue). Modified after Pavlopoulos et al. (2009), with permission from PNAS.

12

(A)

(C)

(D)

(E)

(F)

(G)

(H)

Fig. 4.3.

(B)

(I)

(J)

Bruchdreifachbildung in decapod chelipeds. (A) The possibly oldest account (1671) of this phenomenon showing a right lobster claw (crusher) with duplication of the dactylus and triplication of fixed finger of the propodus. (B) Astacus astacus claw with triple dactylus. (C) The oldest fossil example of a Bruchdreifachbildung. The chela of the Jurassic hermit crab Schobertella simonsenetlangi showing a triple fixed finger. (D) A noble crayfish Astacus astacus with an outer duplication of the right fixed finger. (E) Another specimen of Astacus astacus with a triple fixed finger with the additional elements facing inward. (F) A complicated pattern of horizontal duplications of the fixed finger and the dactylus of the right claw of Astacus astacus. (G) Triple fixed finger of the right claw of the shore crab Carcinus maenas. This image clearly demonstrates the orientation of the multiplied elements. The two outer parts are in the correct orientation; the middle part shows a mirror image (see text). (H) Right claw of a lobster with a horizontal triple fixed finger. As with vertical multiplications, the outer elements are in the right orientation and the middle one is a mirror image. (I) Claw of Cancer pagurus with horizontal duplication of the dactylus. ( J) Lobster claw with the same pattern as the fossil hermit crab in (C). (A) Modified after Berniz (1671). (B) Modified after Rösel von Rosenhof (1755). (C) Photo courtesy of Günther Schweigert. (D–​F) Photos courtesy of Gerhard Scholtz, with two specimen of the Museum für Naturkunde Berlin and one specimen of the Naturalienkabinett Waldenburg. (F) Photos courtesy of Markus Frederich. (H–​J) Photos courtesy of Michel Le Quément from the collection of Charly Le Coadou.

(A)

(B)

(C)

Fig. 4.5.

Patterns of Bruchdreifachbildung in appendages other than chelipeds. (A)  Birgus latro with triplicate second left pereopod. (B) Additional fixed finger in the fourth pereopod of the crayfish Orconectes limosus. (C) Duplication in the antennal scale (scaphocerite) of the freshwater shrimp Paratya curvirostris. (A) Photo courtesy of Jakob Krieger. (B) Photo courtesy of Gerhard Scholtz. (C) Photo courtesy of Stephen Moore.

14

(A)

(B)

(D)

Fig. 4.7.

(C)

(E)

Conjoined twins. (A) A specimen of the freshwater crab Amarinus lacustris with three eyes and a dorsal antenna-​like structure (arrows). This has been interpreted as anterior duplication (duplicitas anterior) of the embryo, with fusion of the median parts such as the eyes and first antennae. (B, C) Two examples of embryonic conjoined twins (duplicitas completa) of the crayfish Cherax destructor (lateral views). (B) Two equal-​size embryos (arrows point to the eyes) are conjoined dorsally, sharing the yolk (y). The embryonic caudal papillae (cp) mark the posterior ends. (C) Two unequal-​size embryos (arrows point to the eyes) are conjoined dorsally, sharing the yolk (y). The left embryo is complete, including the posterior caudal papilla (cp). The right one is smaller and incomplete, with a body rudiment (br) lacking posterior structures. (D) Germ bands of conjoined embryos (duplicitas posterior) of the marbled crayfish Procambarus f. virginalis stained with a fluorescent nuclear marker (ventral view). The left (lle, left embryo’s left eye; lla1, left embryo’s left first antenna; llr, left embryo’s labrum; lcp, left embryo’s caudal papilla) and right (rre, right embryo’s right eye; rra1, right embryo’s right first antenna; rlr, right embryo’s labrum; rcp, right embryo’s caudal papilla) germ bands share a fused eye (fe) and the fused first antenna (fa). All posterior segments are normally developed. (E) Conjoined twins (presumably duplicitas completa) of Lithodes aequispinus (anterior perspective). The two embryos are marked by arrows. (A) Modified after Scholtz et al. (2014), with permission from Elsevier. (B, C) Photos courtesy of Gerhard Scholtz. (D) Photo courtesy of Frederike Alwes and Gerhard Scholtz. (E) Photo courtesy of Bradley G. Stevens.

(A)

(A’)

(B)

(B’)

(B’’) A P

A P S A P S A P S D V D

D V A P

(C)

P

A P

V (D)

(D’)

A

P V

Fig 4.10

(C’’)

(C’)

(E)

(E’)

P P

V

A P

Models explaining the patterns of limb part multiplications and Bruchdreifachbildung. (A, Aʹ) Hans Przibrams models show patterns as a result of regeneration after specific types of injuries. Ventral and dorsal cell states are indicated by red and white areas. According to Przibram, cells tend to regenerate cells of the state that has been removed by the injury. (A) Model explaining the origin and resulting pattern of duplications along the proximodistal axis of a limb. A vertical injury separating the dorsal and ventral cells leads to regeneration of the absent cell states. In this way, a pattern of two-​limb regenerates forms that shows the same dorsoventral orientation. However, such a pattern seems to occur rarely in nature (if at all), because an injury dividing the tip of a limb into two exactly equal-​size lateral halves is an unlikely event. Furthermore, the contact of cells with chirally arranged opposite states in the notch between the regenerated branches is likey to induce an additional tip with a mirror image orientation. Duplications of the rostrum might be a possible example for this (although in a left–​right direction, see Fig. 4.6I). (Aʹ) Model explaining Bruchdreifachbildung. In this case, a lateral injury affecting either the dorsal or ventral cells of the limb and a loss of the tip causes regeneration. As in the previous model, cells of the missing opposite state are regenerated, leading to the specific triplicate pattern, with the two outer regenerates showing the right orientation and the middle structure having a mirror-​image orientation. This is independent of the degree of fusion of the regenerates (see Figs. 4.3–​4.6). (B–​Eʹ) The Boundary Model of Hans Meinhardt explaining the formation of limbs and of multiplication of limb axes based on the boundaries between different cell states. (B–​Bʺ) Formation of arthropod limbs. (B) Schematic representation of segmentation and limb formation. Different colors mark the cell states. Anterior–​dorsal (AD), blue; anterior–​ventral (AV), green; posterior (P), red; separating cell state (S), gray; forming limb, yellow; orientation of the limb, circle with turquoise arrowhead. The limbs form in the area where at least three different cell states meet (AV, AD, P). The topographical relation between ventral and dorsal cells determines the limb orientation as right or left. The gray separating cell state is a requirement to avoid limb formation between the posterior cells of a more anterior segment and the anterior cells of the adjacent segment. These limbs would have a reverse orientation. (Bʹ) Computer model with different cell states (ventral, V; dorsal, D; anterior, A; posterior, P) inducing limb bud growth in the third dimension at the point where these cells stage meet. (Bʺ) The morphological outcome: normal leg. (C–​Cʺ) Explanation of Bruchdreifachbildung with partial fusion of the additional limb. (C) Cell state within a segment with a misplaced patch of posterior cells [color code and abbreviations as in (B)]. The misplaced patch of posterior cells induces additional boundaries between three cell states, resulting in additional limb anlagen. These show the typical orientation, with the middle anlage in a mirror orientation (marked by rotating arrows). (Cʹ) Computerized model of this situation. (Cʺ) The morphological outcome: Bruchdreifachbildung with partly fused additional branches. (D, Dʹ) Similar situation as in (C), but the misplaced patch of posterior cells has a different size and shape. (D) The model. (Dʹ) The morphological outcome: two complete legs with mirror image. (E, Eʹ) Similar situation as in (C), but the misplaced patch of posterior cells has a different size, shape, and position. (E) The model. (Eʹ) The morphological outcome: Bruchdreifachbildung as in (C), but with a complete fusion of the additional limbs. (A, Aʹ) Modified after Przibram (1909). (B–​Eʹ) Modified after Meinhardt (1983, 2009), with permission from Elsevier.

16

(A)

(G)

(B)

(H)

(C) (I)

(D)

(J)

(K)

(E)

(L) (F)

Fig. 6.1.

Larval forms in crustaceans. Comparison with adults (examples). (A–​C) Cirripedia:  Tetraclita squamosal and (C)  T.  formosana. (A)  Nauplius. (B)  Cypris. (C)  Sessile adult. (D–​F) Achelata (Palinuridae). (D)  Palinustus sp. phyllosoma. (E)  Palinustus sp. puerulus. (F)  Palinurus elephas adult. (G–​I) Brachyura:  Dyspanopeus sayi. (G)  Zoea. (H) Megalopa. (I) Adult. ( J–​L) Caridea: Palaemon zariquieyi early ( J) and late (K) zoea. (L) Adult. (A–​C) Photos courtesy of B. Chan (D) Photo courtesy of G. Guerao. (E) Photo courtesy of CSIRO. (F–​I) Photos courtesy of G. Guerao. ( J, K) Photos courtesy of K. Anger. (L) Photo courtesy of B.A. Galán.

1 mm

Young Old

Chalimus 1 Young male

Young female

Chalimus 2 Old male

Old female

Hemianamorphic [metamorphic sensu Atlas of Crustacean Larvae (Martin et al. 2014a)] development patterns exemplified (early anamorphic phase followed by metamorphic change). (A) Developmental stages of Lepeophtheirus salmonis from copepodite to young adult. Notice the strong potential for intrastage growth shown, for example, in the chalimus 2 stage. (A) From Eichner et al. (2015), with permission from Elsevier.

Fig. 6.3.

(A)

Copepodid

18

(A) 3a 4 2

1

3b

8 5

11

(B)

Fig. 6.4.

9

10

7

6

12

13

14

15

(C)

(A) Life cycle of Lepeophtheirus salmonis. Note the presence of anamorphic as well as metamorphic molts. (B, C) Six nauplius stages (N1–​N6) of Eudiaptomus vulgaris (calanid copepod) are followed in (C) by five copepodite stages (C1–​C5). Note that the nauplius stage molts are anamorphic whereas the N6 through C1 molt is metamorphic. (D E) Lepeophtheirus salmonis, nauplius stages 1 (D) and 2 (E). (A) From Piasecki and Avenant-​Oldewage (2008), with permission from Taylor and Francis. (B, C) Modified from Dussart and Defaye (2001), with permission from John Wiley and Sons. (D, E) Modified from Schram (2004), with permission from Cambridge University Press.

Juvenile / adult phase

Metamorphosis

Larval phase

Embryonic phase

Anamorphosis + Metamorphosis = Hemianamorphosis

Metamorphosis

[2]

[1]

Anamorphosis

Hatching

Morphological development

Epimorphosis sensu stricto

Post-Embryonic Moults Fig. 6.7.

Schematic illustration of the principal patterns of postembryonic development in Crustacea. X-axis: Progressing postembryonic molts marked by short vertical lines (left to right). Y-axis: Morphological development (continuous scale, bottom to top). (1) Anamorphosis: (blue line) Hatching occurs with a lower than the adult number of body segments. Initially lacking segments are added during the larval phase. Morphological changes, including formation of appendages, occur typically in small steps at each molt until the juvenile phase is reached. This is typical of taxa with a high number of larval stages (\ \ indicates molts not shown). (2) Hemianamorphosis: (red line) combination of one or more anamorphic phases and one or more metamorphic molts (dramatic morphological changes), followed by epimorphic (sensu stricto) juvenile molts. This is most common pattern in Crustacea. Black dotted vertical lines, transitions between major phases of the life cycle; bold lines, major developmental events. Note that the developmental phase can also be, for example, the zoea phase.

20

(A)

(B)

Fig. 7.5.

The microplastics ingested by crustacean larvae are visualized under fluorescence microscopy (fluorescing yellow-​ green in guts) following laboratory exposure of the larvae to fluorescent polystyrene beads. (A) A porcellanid larva containing 30.6-​μm polystyrene beads (lateral view). (B) A brachyuran crab zoea larva containing 20.6-​μm polystyrene beads (lateral view). Modified from Cole et al. (2013), with permission from ACS, © 2013 American Chemical Society. Photos courtesy of Dr. Cole.

(A)

(D)

Exhalant siphon

0.01

Inhalant siphon

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Rectum

–0.01

Stigmata

Test

Branchial basket Intestine

i –0.01

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0.01

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Esophagus Stomach Flow Food (B)

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y position (mm)

–0.01 ii –0.01

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(C) Water flow Food particles Buccal cavity Gill arch Gill filament Water flow

Water flow

Fluid speed (m/sec)

0.01 0 –0.01 vi x position (mm)

Fig. 13.2.

Predator feeding modes. (A) Diagram of a current-​feeding ascidian (Pyura stolonifera) predator. Modified from Knott et al. (2004), with permission from the Marine Biological Laboratory, Woods Hole, MA. (B) Grasping scyphozoan (Eutima gracilis) predator (12.5 mm across) eating a copepod. Modified from Lebour (1923), with permission from Cambridge University Press. (C)  Fluid velocities produced by a suction-​feeding bluegill (Lepomis macrochirus), showing (i) opening, (ii) prey entering, (iii) peak gape, and (iv) closing during the strike. Modified from Day et al. (2005), with permission from the Company of Biologists, Ltd. (D) Diagram of a filter-​feeding fish. Modified from Paig-​Tran et al. 2011, with permission from the Company of Biologists, Ltd.

MB PB PC PV 32 31

MB

PB PC

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32

31 PV

PC PB

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PR PA CM

30 m

CBHB

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32

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PC PB

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DVM

CBHB

10–4

10–3

10–2

10–1

larval density/(km2)

Numerical model of larval dispersion and connectivity for larvae (log10 density scale) regulating depth in four ways during the peak upwelling season (spring) in the California Current System: maintaining a depth of 5 m, maintaining a depth of 30 m, undertaking an ontogenetic vertical migration (OVM) from 5 to 30 m deep, and undertaking a diel vertical migration (DVM) from 5 to 30 m deep. Dispersion after 30 days for larvae released near Bodega Head, California (•), and connectivity after 30 to 60 days for larvae released and recruiting at sites from Palos Verdes, California, to Heceta Bank, Oregon. Diagonal line in connectivity matrices represents local recruitment, with intensity above it representing poleward dispersal and intensity below it representing equatorward dispersal. PV, Palos Verdes; PC, Point Conception; PB, Point Buchon; (MB), Monterey Bay; PR, Point Reyes; PA, Point Arena; CM, Cape Mendocino; CB, Cape Blanco; HB, Heceta Bank. Modified from Drake et al. (2013).

Fig. 14.3.

PR

PR

CB HB

PA

PA

PR PA CM

CM

CM

MB

HB CB

HB CB

PC PB

47 CR

47 CR

PV

5m

Connectivity

Dispersion

2

(A)

Fig. 15.5.

(B)

Model results: dispersal distance for two larval types. Maps showing the average dispersal distance in kilometers for each grid cell at a 0- to 100- m depth, where the trajectories were released for two different larval types: (1) drifting at the surface (0–2 m) and (2) below the pycnocline (24–26 m), with a 30- day pelagic larval duration (PLD). Marine protected areas in the Kattegat- Skagerrak- Belt Sea area are marked on the maps with black lines. From Jonsson et al. 2016, with permission from John Wiley and Sons.

24

Fig. 15.6.

(A)

(B)

(C)

(D)

Model results: source– sink analyses for two larval types in eastern Skagerrak. (A) and (B) show the distri­bution of larval sources for eastern Skagerrak (northwest coast of Sweden), where the larvae that settled in eastern Skagerrak were released. (C) and (D) show the larval sinks for eastern Skagerrak, were larvae released in eastern Skagerrak have settled, for two types of larvae: trajectories drifting at the surface (0– 2 m) and those below the pycnocline (24– 26 m) during a 30- day larval phase. All trajectories where released and settled at 0-100- m depth. Darker color denotes higher frequency of trajectories. White denotes zero trajectories. Modified from Moksnes et al. (2014b).

1 FROM A SINGLE CELL TO SEGMENTAL STRUCTURES: CRUSTACEAN EMBRYOLOGY

Gerhard Scholtz

Abstract Beginning with Aristotle 2400 years ago, research on crustacean embryology has a long tradition. Rathke’s 1829 landmark study on the noble crayfish initiated modern approaches. Crustaceans in general—​and most of their large taxa—​show a great diversity in all stages of their developmental pathways from the zygote up to the adult animal. This chapter describes the various modes of cleavage, gastrulation, germ band formation, and segmentation found in crustacean taxa. Cleavage is either total, partial, or mixed. Total cleavage can be indeterminate, without predictable cell lineage; or determinate, with a stereotyped cell division pattern. Gastrulation modes can also vary to a high degree. One finds invagination, epiboly, immigration, delamination, and a mix of some of these. Likewise, the stages of germ layer separation and the number of cells that initiate gastrulation differ. In yolk-​rich eggs, a germ disk forms at the future ventral side of the embryo, and the axes and orientation of the germ are recognizable. Through elongation in the anteroposterior direction by a posterior growth zone and intercalary cell divisions, the germ disk is transformed into the germ band. As a result of a unique, stereotyped cell division pattern in the germ band of malacostracans, germ band growth and the segmentation process up to the differentiation of neuronal precursors and early limb anlagen can be analyzed at the level of individual cells. Recent morphological and molecular techniques allow a very detailed spatiotemporal resolution of developmental processes and they offer new perspectives on long-​standing morphological questions.

INTRODUCTION: THE DIVERSITY OF CRUSTACEAN ONTOGENIES The great diversity of adult crustacean forms, body organizations, and lifestyles finds its correspondence in a comparable variety of ontogenetic pathways. Crustaceans are unmatched by other Developmental Biology and Larval Ecology. Edited by Klaus Anger, Steffen Harzsch, and Martin Thiel. © 2020 Oxford University Press. Published 2020 by Oxford University Press.

1

2

2

Developmental Biology and Larval Ecology arthropod groups with regard to their manifold cleavage and gastrulation patterns, segmentation processes, and larval types and development. This fascinating ontogenetic diversity is even greater if recent views on crustacean phylogeny are taken into account. According to these views, crustaceans are paraphyletic with Remipedia, either alone or together with Cephalocarida, as a sister taxon to Hexapoda (Regier et al. 2010, von Reumont et al. 2012, Schwentner et al. 2017, see Chapter 5 in Volume 8). Hence, insects are deeply nested within a group that was traditionally called Crustacea but is now named Pancrustacea or Tetraconata (Zrzavý and Štys 1997, Dohle 2001). Yet, this ontogenetic diversity and the new phylogenetic aspects make any general statements or a reconstruction of a ground pattern very difficult. The following account is restricted to crustaceans in the traditional sense. However, when necessary, reference is also made to hexapods. Our knowledge of the ontogeny of crustaceans varies widely across the different taxa. For instance, there are no data at all about the early development of Remipedia. Only the larval development of one species has been described to some extent (Koenemann et  al. 2009). Similarly, besides some data on the postembryonic development, nothing is known about the embryology of Mystacocarida and Tantulocarida (Huys et al. 2014, Olesen and Haug 2014). Until recently, knowledge on the ontogeny of Cephalocarida has also been restricted to an external description of the eggs and the larvae (Olesen et al. 2014b). Yet, now there are also some first results on nervous system development (Stegner and Richter 2015). Likewise, one recent description of branchiuran embryology was published by Banerjee et al. (2015), which complements studies on more advanced stages (Kaji et al. 2011). The putative sister group of Branchiura, the Pentastomida (see Møller et al. 2008, Regier et al. 2010), are also covered by an embryological study (Osche 1963). There are several investigations that deal with the embryonic and larval development of Thecostraca (see Anderson 1973, Deutsch et  al. 2004, Scholtz et  al. 2009b, Semmler et al. 2009, Ponomarenko 2014, Høeg et al. 2014, 2015). Given the great numbers of species, Copepoda and Ostracoda are relatively poorly studied with respect to their embryology. Yet, at least some detailed studies exist on the early development of these two groups (e.g., Grobben 1881, Schimkewitsch 1896, McClendon 1907, Müller-​Calé 1913, Fuchs 1914, Witschi 1934, Weygoldt 1960, Kohler 1976, Wakayama 2007, Hein et al. 2019, Koyama and Rivera 2019, Loose and Scholtz 2019). In the case of Branchiopoda, the situation is heterogeneous. For Notostraca, Laevicaudata, and Spinicaudata, there are no proper studies on the early embryonic development, but there are many on the larvae (e.g., Olesen 2004, Pabst and Scholtz 2009, Fritsch et al. 2013). In Anostraca, our knowledge about early embryonic development is largely limited to Artemia salina (Benesch 1969). However, a number of investigations dealing with limb formation, segmentation, and neurogenesis have been carried out, including other anostracan species (e.g., Fränsemeier 1939, Anderson 1967, Freeman 1986, Panganiban et al. 1995, Manzanares et al. 1996, Olesen 2004, Williams 2004, Williams et al. 2012). Cladocera is relatively well covered by a number of embryological studies, which include all of the larger groups: Haplopoda (Samter 1900, Gerberding 1997), Anomopoda (Grobben 1879, Mittmann et al. 2014), Onychopoda (Kühn 1913, Alwes and Scholtz 2014), and Ctenopoda (von Baldass 1937). Daphnia magna has been established as a model organism for molecular developmental analyses (Sagawa et al. 2005, Ungerer et al. 2011, 2012, Kato et al. 2012, Mittmann et al. 2014). A similar situation applies for the Malacostraca, in which embryological data have been collected for all major taxa—​namely, Leptostraca (Manton 1934, Olesen and Walossek 2000, Pabst and Scholtz 2009); Stomatopoda (Shiino 1942); Decapoda (e.g., Brooks 1882, Reichenbach 1886, Weldon 1892, Sollaud 1923, Ooishi 1959, 1960, Scheidegger 1976, Zilch 1978, Biffis et al. 2009, Klann and Scholtz 2014, Harzsch et al. 2015, Hertzler 2015, Alves et al. 2019, Ma et al. 2019); Syncarida (Hickmann 1937); Euphausiacea (Taube 1909, Alwes and Scholtz 2004, Jia et  al. 2014, Ambriz-​Arreola et  al. 2015); Thermosbaenacea (Zilch 1974); and Mysidacea, Amphipoda, Tanaidacea, Cumacea, and Isopoda among the Peracarida (e.g., van Beneden and Bessels 1868, Ulianin 1881, Bergh 1893, 1894, McMurrich 1895, Langenbeck 1898, Manton 1928, Weygoldt 1958, Scholl 1963, Strömberg 1965, 1971,



From a Single Cell to Segmental Structures

Petriconi 1968, Dohle 1970, 1972, 1976, Mergault and Charniaux-​Cotton 1973, Scholtz 1984, 1990, Gerberding et al 2002, Wolff and Scholtz 2002, Wolff 2009). This list is by no means complete but it is evident that there are only a few studies dealing with Leptostraca, Stomatopoda, Syncarida, Thermosbaenacea, and Euphausiacea. In contrast to this, Peracarida [with the exception of the smaller taxa such as Spelaeogriphacea and Mictacea (see Olesen et  al. 2014a)] and Decapoda have been examined relatively comprehensively, including some recent molecular developmental studies. Hence, at first sight, it looks as though we are pretty well informed about the embryonic development of crustaceans in general. However, it must again be stressed that for many groups of crustaceans, there are only a few detailed studies and that many of the investigations originate from the 19th or early 20th centuries. This does not mean there are not any outstanding studies from this period, with evidence and results that are still valid today. Yet, one should be aware that the technical and methodical limitations of those times make some results problematic. This concerns primarily the allocation of embryonic cells to a germ layer or to a specific cell fate. Interpolations and extrapolations were often made without real evidence, and conclusions were drawn that can actually only be proven using the techniques available today, such as single-​cell labeling, confocal laser scanning microscopy, four-​dimensional microscopy, and the use of molecular markers. Furthermore, the authors at that time did not yet possess the methodology and logic of phylogenetic systematics to interpret their data comparatively in a phylogenetic and evolutionary context. In summary, it can be asserted that there is a significant need for more comparative embryological studies in crustaceans.

A SHORT HISTORICAL ACCOUNT OF CRUSTACEAN EMBRYOLOGY Aristotle was the first to mention crustacean eggs in a scientific context. In his descriptions of crustacean anatomy and development in his books Historia Animalium, De Partibus Animalium, and De Generatione Animalium (ca. 340 BC) [Aristotle (ca. 340 BC) with English translations by Cresswell, Ogle, and Peck], he reported on testes, ovaries, and the eggs of the spiny lobster and other decapods. Aristotle considered them imperfect eggs because they enlarge during the first phase of development and they lead to larvae. He described the eggs of spiny lobsters as a grainy mass attached to the pleopods. The 18th and early 19th centuries saw a number of cursory descriptions of external aspects of the embryology of freshwater crayfish, copepods, and various branchiopods (Rösel von Rosenhof 1755, Cavolini 1792, Prévost 1803, Jurine 1820). These descriptions were embedded in comprehensive and detailed natural history accounts. The first detailed treatment of crustacean embryology dates back to 1829, when Heinrich Rathke published his article on the development of the freshwater crayfish Astacus astacus. Rathke’s investigation is part of the earliest set of embryological studies on invertebrates that began with Johan Herold’s account of butterflies (1815) and spiders (1824) (see Wellmann 2013). Rathke used a table structure to illustrate the morphological changes of the embryo over time, which Herold and he had adopted from the earlier vertebrate embryologists and is still used in recent developmental publications [see Wellmann (2010), Hopwood (2007), and Scholtz (2014) for historical accounts] (Fig. 1.1). Some descriptions of Rathke’s studies were very detailed. For instance, he was the first to detect the yolk pyramids in crayfish embryos. Rathke continued with descriptions of the embryology of several isopod species and other crustaceans (e.g., Rathke 1837). These early studies dealt mainly with the outer shape of the embryos and the aspects of adult form generation. The understanding of the role of cleavage, cell divisions, and germ layers was still in its infancy, in particular for invertebrates. For instance, during the mid 19th century, scientists still

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Fig. 1.1. The arrangement of the developmental stages. A plate of the pioneer work of Heinrich Rathke (1829) showing stages of the development of the freshwater crayfish Astacus astacus. Modified after Rathke (1829).



From a Single Cell to Segmental Structures

argued whether crustaceans showed a proper cleavage, as found in other animals (see Leydig 1848). Soon after, it was clear that this was the case, as exemplified by the observation of total cleavage in rhizocephalans by Müller (1862). Nevertheless, most of the studies of this time are mainly of historical interest. The role of the knowledge of animal ontogeny for systematics became evident as early as the 1830s, when Thompson detected the nauplius of barnacles. This finding led to the conclusion that they are indeed crustaceans (Thompson 1830), and it ended the long discussion about the systematic position of these enigmatic animals that were interpreted as embryonic geese or mollusks (Scholtz 2008, Buckeridge and Watts 2012). Darwin (1859) himself stressed that ontogeny should be included in the reconstruction of genealogical evolutionary relationships, because early stages often display more and more obvious resemblances between taxa than the adult animals. Five years after Darwin’s epochal work, Fritz Müller published a little book dedicated to Darwin, titled Für Darwin, in which he refined the old idea of recapitulation in a strictly evolutionary context (Müller 1864). This approach was based on the detection of the nauplius in a dendrobranchiate shrimp, which led Müller to the idea that the ancestor of crustaceans as a whole was a nauplius (Müller 1864). Haeckel adopted, extended, and propagated Müllers views, and formulated the still-​popular law of recapitulation: The ontogeny is a quick recapitulation of phylogeny (Haeckel 1866). The period from the 1870s to the early decades of the 20th century represents the heyday of comparative embryology in general (Bowler 1996), and that of crustaceans in particular. This boost was triggered mainly by two developments. First, the rise of evolutionary theory in combination with the concepts of germ layers and recapitulation opened a fruitful theoretical background for the interpretation of morphological structures and their ontogenetic generation. Second, technical improvements such as the invention of the microtome, histological staining methods, and better microscopes led to a dramatically enhanced resolution of morphological structures at the levels of cells, tissues, germ layers, and organs. During the golden age of evolutionary embryology, numerous studies were published that dealt with cleavage patterns, germ layer formation, and morphogenesis of organs of representative species of most crustacean groups. In a seminal study on the early development of the cladoceran Moina rectirostris, Grobben (1879) pioneered the combination of cleavage and the formation of germ layers. Following this approach, the determinate cleavage patterns of cirripeds, copepods, decapods, euphausiaceans, amphipods, cladocerans, and ostracods were described by several authors (e.g., Grobben 1881, Brooks 1882, Langenbeck 1898, Bigelow 1902, Taube 1909, Kühn 1913, Müller-​Calé 1913, Fuchs 1914). Because the prevailing theory at that time was that arthropods were closely related to Annelida based on the shared body segmentation, crustacean cleavage was often interpreted with respect to spiral cleavage (e.g., Taube 1909, Fuchs 1914). Additional important steps in crustacean embryology were noted by Reichenbach (1886), who detected the special stem cells of the growth zone of malacostracans: the ectodermal teloblasts. Patten (1890) described similar cells in the mesoderm: the mesoteloblasts. Bergh (1893, 1894) and McMurrich (1895) were the first to resolve aspects of the regular cleavage pattern in the malacostracan germ band and reported the existence of neural stem cells—​the neuroblasts comparable to those that were described for hexapods a few years earlier (Wheeler 1891). By the end of the 19th century, comparative methods of morphology and embryology lost their leading role in evolutionary biology and interest shifted to experimental approaches. In particular, developmental studies turned more to the direction of uncovering developmental mechanisms by experimental manipulation of embryos. This physiological approach of the Entwicklungsmechanik, as propagated by Wilhelm Roux (1894, 1907), initiated a new field that had dominated developmental biology since then (Mocek 1998). The modern version of this approach is molecular developmental biology in all its facets.

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Developmental Biology and Larval Ecology Interestingly enough, none of the determined cleavage patterns of crustaceans reached iconic status like spiral cleavage (Henry 2014) or the early development of nematodes (Schnabel 1997). Likewise, no crustacean gained a role as a model organism like Drosophila melanogaster (Lawrence 1992) or Parascaris sp. (later replaced by the nematode Caenorhabditis elegans) (Moritz and Sander 1996). This might be due to several reasons. One is the diversity found in crustaceans, which renders the search for a “typical” representative problematic. For instance, the patchy distribution of cleavage modes such as superficial versus total and determinate versus indeterminate cleavage makes it difficult to arrive at a safe conclusion concerning ancestral crustacean cleavage. In addition, any conclusions or generalizations are hampered by the largely unresolved phylogenetic relationships among crustaceans, notwithstanding the recent view of crustaceans being paraphyletic with respect to hexapods (see Chapter 5 in Volume 8). Other reasons might be found in the fact that the maintenance of brood stocks of most crustaceans is complex and laborious. Finally, yet importantly, the delicacy of most eggs and embryos renders manipulations problematic, and in many cases there is just a seasonal supply with eggs and a long generation cycle. For similar reasons, crustaceans were rarely used for experimental approaches to uncover developmental mechanisms. Yet, some centrifugation experiments were carried out on copepod and cladoceran eggs (Spooner 1911, Kaudewitz 1950). Jacobs (1925) ablated single cells of the two-​and four-​cell stages of a copepod, and Kajishima (1951, 1952a,b) ablated parts of isopod, cirriped, and penaeid embryos. Nevertheless, most of the 20th century saw some very detailed comparative studies on crustacean embryology (e.g., Manton 1928, 1934, von Baldass 1937, 1941, Shiino 1942, 1950, Weygoldt 1958, 1961, Ooishi 1959, 1960, Scholl 1963, Anderson 1969, Dohle 1970, 1972, 1976, Fioroni 1970a, Zilch 1974, 1978, Scheidegger 1976, Scholtz 1984, 1990). These investigations inspired a number of textbooks (Siewing 1969, Anderson 1973, Schwartz 1973, Fioroni 1987). However, most of these studies had little impact beyond the specialists’ field. This changed with the rise of evolutionary developmental biology (EvoDevo) during the 1980s. The comparative approach that is central to this discipline revived the interest in crustacean embryology. The morphological diversity of crustaceans and the new view on their phylogenetic relationships as close relatives of Hexapoda put them into the focus of arthropod evolution. In particular, the possibility of using malacostracan species to combine detailed cell lineage studies with gene expression offered a new dimension and resolution unmatched by other arthropod embryos (e.g., Dohle et al. 2004, Serano et al. 2016, Wolff et al. 2018). Recent morphological and molecular techniques such as improved methods of cell ablation and isolation, confocal laser-​scanning microscopy with computer-​aided three-​dimensional reconstructions, single-​cell labeling, four-​dimensional microscopy, light-​sheet microscopy, cross-​ reacting antibodies, in situ hybridization in nonmodel organisms, and genetic manipulations (RNA interference, transgenesis, and CRISPR/​Cas9-​based somatic mutagenesis) led to a spatial and temporal resolution of crustacean development that was previously unknown (Stamataki and Pavlopoulos 2016). These studies help to clarify developmental mechanisms, but they also offer the possibility of arriving at clearer conclusions about the homology of crustacean body parts and the evolution of the various limb types and tagmosis patterns. Questions such as the nature of appendages, the evolutionary transformation of pereopods to maxillipeds, segmentation of the head, the relationships between the limbless abdomen of non-malacostracan and the malacostracan pleon, and the origin of hexapod wings can be addressed based on a new set of data (e.g., Averof and Akam 1995, Panganiban et  al. 1995, Scholtz 1995, 2016, Scholtz et  al. 1998, 2009b, Averof and Cohen 1997, Abzhanov and Kaufman 2004, Deutsch et al. 2004, Pavlopoulos et al. 2009, Martin et al. 2016, Fritsch and Richter 2017, Wolff et al. 2018).



From a Single Cell to Segmental Structures

CRUSTACEAN EMBRYOLOGY Cleavage Total, Partial, and Mixed Cleavage Modes Cleavage can be conceptualized as the process that subdivides the zygote into smaller compartments. This compartmentalization (in combination with a graded distribution of morphogens) is the prerequisite for subsequent differentiation and fate determination of embryonic regions. Cleavage begins with the first mitotic division of the zygote and ends with the blastula or blastoderm stage. However, the latter is frequently not expressed in a typical manner as a result of an overlap with early gastrulation. Crustaceans show many of the cleavage modes listed in textbooks. There are yolky eggs and those with almost no yolk (Figs. 1.2–​1.7). Sometimes the animal vegetal axis of the egg is indicated by the occurrence of polar bodies, but frequently it is only recognizable with the beginning of gastrulation. One finds total cleavage, partial cleavage, and mixed cleavages, which combine both modes (Fig. 1.6). Total cleavage is either equal, with blastomeres of similar size; or unequal, characterized by the occurrence of large macromeres and small micromeres (Fig. 1.2). Furthermore, it can be determinate with stereotyped cleavage patterns and cell fates or indeterminate (Fig. 1.2). These cleavage modes are often correlated with size and yolk content of the eggs. Total cleavage occurs in cases with small eggs showing a little amount of yolk; partial cleavage is found in large and yolk-​rich eggs (Figs. 1.2 and 1.3). Yet although this is frequently the case, this correlation is not universal. Large and yolky eggs of amphipod species undergo total cleavage (Scholtz and Wolff 2002) (Fig.1.2), whereas some tiny eggs of cladocerans such as the haplopod Leptodora kindtii display partial cleavage (Samter 1900) (Fig. 1.3). In these cases, the inherited pattern is maintained despite the change of egg size and yolk content (Scholtz and Wolff 2002). Total cleavage is characterized by a complete separation of the blastomeres by cell membranes. In addition, often a blastula stage with a single cell layer surrounding a central liquid-​filled space, the blastocoel, is formed. This cleavage mode occurs among Anostraca (Benesch 1969), Thecostraca (Scholtz et al. 2009b), Ostracoda (Müller-​Calé 1913), Copepoda (Fuchs 1914, Witschi 1934, Loose and Scholtz 2019), Amphipoda (Scholtz and Wolff 2002), Euphausiacea (Alwes and Scholtz 2004), Dendrobranchiata (Hertzler 2015), Anaspidacea (Hickman 1937), and the parasitic isopod Hemioniscus balani (Caullery and Mesnil 1901). Thus, the size of the blastomeres is more or less alike (equal cleavage) in ostracods, free-​living copepods, dendrobranchiates, euphausiaceans, and anaspidaceans (Figs. 1.2 and 1.5). In amphipods, however, a clear discrimination between micromeres and macromeres (unequal cleavage) is possible, beginning with the eight-​cell stage (Gerberding et al. 2002, Wolff and Scholtz 2002) (Figs. 1.2 and 1.5). Thecostraca show an extreme unequal cleavage. The first cleavage division of Thoracica, Acrothoracica, and Ascothoracida results in a large blastomere containing most of the yolk and an almost yolkless micromere (Scholtz et al. 2009b, Ponomarenko 2014) (Fig. 1.2). Further asymmetric divisions of the macromere and symmetric divisions of the micromeres produce more micromeres (Anderson 1973, Ponomarenko 2014). Partial cleavage means that the products of early cleavage divisions are not separated by membranes. In the case of superficial cleavage, only nuclei surrounded by yolk-​free cytoplasm, the so-​called energids, divide and are situated within the yolk mass (Fig. 1.3). After a certain round of cleavage divisions, the energids migrate to the egg periphery and form a cellular blastoderm (Fig. 1.3). This superficial cleavage, as exemplified by the development of the model organism Drosophila melanogaster (see Lawrence 1992), is also frequently present in crustaceans—​for instance in cladocerans (Samter 1900), branchiurans (Banerjee et al. 2015), decapods (Scheidegger

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Fig. 1.2. Total and mixed cleavage of various crustaceans with a stereotyped cell division pattern. (A) The amphipod Cryptorchestia garbinii. (B) The euphausiid Meganyctiphanes norvegica. (C) The dendrobranchiate decapod Sicyonia ingentis. (D) Dorsal aspect of the cirriped Austrominius modestus. (Dʹ) Ventral aspect. (E) Vegetal aspect the cladoceran Bythotrephes longimanus. (Eʹ) Lateral aspect. Sister cells are connected with a black line. Except for the amphipod C. garbinii yellow marks the A quadrant; red, the B quadrant; green, the C quadrant; and blue, the D quadrant, indicating putative homologies. The nomenclature of individual cells is only added in B. longimanus (en, cells forming the endoderm, g, the primordial germ cells). The cleavage of Amphipoda is so different from the other crustaceans that the coloring is chosen arbitrarily. The arrows point to the gastrulation center. In malacostracans, it occurs at the end of a cell band (blue blastomeres meeting the center of the yellow–​green band); in non-malacostracans, in the center of a cell band (contact zone of the blue and red blastomeres). Each column shows, from top to bottom, the four-​cell stage, the eight-​cell stage, the 16-​cell stage, and the 32 cell-​stage (the latter is not, in all cases, complete). B. longimanus undergoes a mixed cleavage in which the early stages are characterized by a central yolk mass. C. garbinii, A. modestus, and B. longimanus display unequal cleavages (micromeres and macromeres), whereas those of M. norvegica and S. ingentis are equal. The two interlocking cell bands (tennis ball pattern) is best seen in the euphausiacean and the decapod. In the cirriped, it is slightly transformed in the B and D quadrants; in the cladoceran, the contact zone (cross-​furrow) of the B and D quadrants is found at the vegetal pole, and of the A and C quadrants at the animal pole, as in the cirriped, despite an arrangement lacking the characteristic cell bands. (A) Modified after Scholtz and Wolff (2013, Fig. 4.4) and Scholtz and Wolff (2002, Fig. 4c). (B) Modified after Alwes and Scholtz (2004, Fig. 5). (C) Modified after Hertzler and Clark (1992, Fig. 6). (D, Dʹ) Modified after Ponomarenko (2014, Fig. 3.3.1). (E, Eʹ) Modified after Alwes and Scholtz (2014, Fig. 7). See color version of this figure in the centerfold.

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Fig. 1.3. Superficial cleavage. (A–​F) From the two nucleus stage to the 32-cell stage of the cleavage of the isopod Idotea. The spindle axes leading to the four-​nuclei stage show a 90-​degree angle to each other. The nuclei reach the egg’s surface between the 16-​and the 32-​cell stages. Cell boundaries are formed at the 32-​cell stage. (G) Four-​ nuclei stage of the isopod Porcellio scaber with the same arrangement of the nuclei (dark areas) within the yolk mass as in (B). (H–​K) Indications of cell fate determination in superficial cleavage. (H) Transition between the 32-​and 64-​cell stages of the cumacean Diastylis rathkei. Two cells show a retarded division and are the first to form cell boundaries. These probably mark the future gastrulation center. (I–​K) Sixteen-​to 64-​cell stages of the cladoceran Leptodora kindtii. In this case, superficial cleavage forms the pattern of two interlocking bands comparable with that of some total cleavages (see Fig. 1.2). (A–​F) Modified after Strömberg (1965, Fig.  1). (G) Modified after Scholtz and Wolff (2013, Fig. 4.5). (H) Modified after Dohle (1970, Fig. 5). (I–​K) Modified after Alwes and Scholtz (2014, Fig. 10). See color version of this figure in the centerfold.

1976), leptostracans (Manton 1934), stomatopods (Shiino 1942), and most peracarids (Manton 1928, Scholl 1963, Dohle 1970, Strömberg 1965, Scholtz and Wolff 2013). There are several examples in which partial cleavage is combined with superficial cleavage furrows (e.g., Weygoldt 1961, Scheidegger 1976). This leads to an outer appearance of the eggs that is comparable to that of total cleavage. Yet, the furrows do not entirely penetrate the yolk; hence, there is always a large, central undivided yolk mass (Weygoldt 1961, Strömberg 1971, Scheidegger 1976). This cleavage is, nevertheless, of the superficial mode and has to be discriminated from mixed cleavage, as described later. Some groups such as freshwater crayfish (Rathke 1829, Reichenbach 1886), lobsters (Herrick 1895), and stomatopods (Shiino 1942)  with yolk-​rich eggs and superficial cleavage differentiate so-​called yolk pyramids (Fioroni 1970a) (Fig. 1.4). Yolk pyramids occur with the final stages of cleavage, in particular during blastoderm formation, and subdivide the central yolk mass into smaller compartments (Reichenbach 1886). The formation of the yolk pyramids is not fully understood. In freshwater crayfish, they form by propagating from the egg periphery toward its center (Reichenbach 1886, Nobis 2012) (Fig. 1.4). This way, the yolk is incorporated into tapering hexagonal compartments. In the egg’s center, an undivided central yolk mass remains. It is not clear whether the yolk pyramids are separated by proper cell membranes. There are some hints that this is the case, but further studies using transmission electron microscopy are required (Nobis 2012) (Fig. 1.4). Later, the walls of the yolk pyramids disintegrate; a large central yolk mass is formed underneath the blastoderm when



From a Single Cell to Segmental Structures (A)

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Fig. 1.4. Yolk pyramids in the freshwater crayfish Procambarus f. virginalis. Except for (D), which is a fluorescent micrograph, all images were taken with a scanning electron microscope. (A) External view in the early blastoderm stage. (B)  Fracture showing the early yolk pyramids (yp) growing inward from the blastoderm toward the central yolk mass (y). (C) Fracture displaying slightly advanced growth of yolk pyramids (yp). These assumed their characteristic hexagonal shape. (D) Treatment with phalloidin fluorescent staining indicates the membranous nature (arrows) of the boundaries of the yolk pyramids (yp). The hexagonal patterns (upper arrow) mark the transition to the central yolk mass (y). (E) The maximal extension of yolk pyramids (yp) with a small central yolk mass (y). (F) Largely degenerated yolk pyramids (yp) at the stage of initial gastrulation. The central yolk (y) is again large. (A) Modified after Alwes and Scholtz (2006, Fig. 2a). (B–​F) Modified after Nobis (2012 Figs. 6a, 7a, 8b, 10a and 14a).

gastrulation starts. In ontogenies in which the forming gut encloses large amounts of yolk, secondary and sometimes tertiary yolk pyramids have been described (Fioroni 1970a) (Fig. 1.7K). Again, these secondary yolk pyramids subdivide the yolk into smaller compartments, but their formation and their nature are even less obvious. Primary yolk pyramids have sometimes been interpreted as remains of a former total cleavage, but this is not clear. Mixed cleavage (Fioroni 1970a, Scholtz and Wolff 2013) as is found in most Cladocera (Alwes and Scholtz 2014), parasitic Copepoda (McClendon 1907, Kohler 1976), Thermosbaenacea (Zilch 1974), parasitic Isopoda, and many pleocyemate Decapoda (Müller 1984, Klann and Scholtz 2014, Romero-​Carvajal et al. 2018) shows a combination of elements of total and partial cleavage modes (Figs. 1.2E, Eʹ and 1.6). Sometimes the first cleavage divisions are superficial, without membranes or with membranes, reaching only partially into the yolk (Alwes and Scholtz 2014). Later, the divisions switch to total cleavage or vice versa (Weldon 1892, Strömberg 1971, Zilch 1974, Müller 1984). Like total cleavage, mixed cleavage can follow an equal or unequal pattern, with different blastomere sizes. Examples of a mixed cleavage with equal-​size blastomeres are found among decapods and anomopodan cladocerans (von Baldass 1941, Klann and Scholtz 2014); unequal cleavage is exemplified by onychopodan cladocerans (Kühn 1913, Alwes and Scholtz 2014) (Fig. 1.2E, Eʹ). An extremely unequal mixed cleavage pattern occurs in parasitic copepods, in which the first cleavage leads to a large blastomere containing all the yolk and a small micromere (Schimkewitsch 1896, McClendon 1907, Kohler 1976).

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Fig. 1.5. Cell lineage studies. (A, B) The amphipod Cryptorchestia garbinii. (C, D) The cirriped Austrominius modestus. (E) The decapod Sicyonia ingentis. (A) Live images of the cleavage from the zygote to early gastrulation. The image of the 16-​cell stage in the center is a scanning electron microscope micrograph. (B) The result of the single-​cell application of the fluorescent dye DiI to two micromeres (shown in red in the inset). The entire postnaupliar mesoderm, including the mesoteloblasts plus the endodermal midgut glands, show the staining. (C) Ventral (left) and dorsal (right) aspects of the transition of the 16-​to 32-​cell stage (Sytox green marker of cell nuclei). (D) Maps of the contributions of the quadrants to the external ventral (left) and dorsal (right) sides and internal organs (nervous system, musculature, intestine) of the nauplius larva. (E) Cell lineage markings of the blastomeres of the four-​cell stage and their outcome (dotted areas) in the nauplius larva. The upper row in each box shows the ventral side; the lower row, the dorsal side. Mirror images are the results of the two chiral (mirror image) variants of eggs. (A) Modified after Scholtz and Wolff (2002, Fig. 1). (B) Modified after Wolff and Scholtz (2002, Fig. 12). (C, D) Modified after Ponomarenko (2014, Figs. 3.3.6 and 3.4.1). (E) Modified after Hertzler and Clark (1992, Fig. 6) and Hertzler et al. (1994, Fig. 4). See color version of this figure in the centerfold.



From a Single Cell to Segmental Structures

Determinate Cleavage The total cleavage of Anostraca, for instance, does not follow a predictable pattern of cell divisions and cell arrangements (Benesch 1969). Moreover, the partial cleavage with superficial furrows of the hermit crab Pagurus prideaux and the mixed cleavages of some caridean shrimp and parasitic isopods show a great variability in the arrangement of the early energids and blastomeres (Gorham 1895, Strömberg 1971, Scheidegger 1976, Klann and Scholtz 2014). Yet, in a number of crustacean taxa, the total and mixed cleavage modes are determinate. In other words, they show a stereotyped cell division pattern with respect to the topographical relation and size of blastomeres, and the spindle directions of blastomere mitoses. Furthermore, they reveal predictable cell fates. The cooccurrence of a distinct cell division pattern and determined cell fates is called cell lineage. Prominent examples of cleavage modes with cell lineages are the spiral cleavage of mollusks, annelids, and a number of other taxa (Henry 2014), and the cleavage of nematodes such as Caenorhabditis elegans (Schnabel 1997). A determinate cleavage is found in representatives of cirripeds (Bigelow 1902, Anderson 1969, Ponomarenko 2014), copepods (Fuchs 1914), and cladocerans (Kühn 1913, von Baldass 1937, 1941, Alwes and Scholtz 2014). Furthermore, it occurs in euphausids (Taube 1909, Alwes and Scholtz 2004), dendrobranchiates (Zilch 1978, Hertzler and Clark 1992, Hertzler 2015), and amphipods (Gerberding et  al. 2002, Wolff and Scholtz 2002)  among malacostracans (Fig. 1.2). However, these cleavage patterns are largely taxon specific and it is difficult to extract general aspects. This is exemplified by a comparison of the cleavage pattern of an amphipod, a euphausiacean, and a dendrobranchiate decapod (see Scholtz and Wolff 2013) (Fig. 1.2A–​C). The patterns of the euphausiacean and the dendrobranchiate decapod correspond to each other in many details, such as the round shape of the egg, and the size and arrangement of the blastomeres, which form two interlocking cell bands. Furthermore, the gastrulation centers of both species, including the endoderm and the prospective germ cells, are situated in the same area and are generated by a corresponding cell lineage. In summary, there is evidence that these cleavage patterns are homologous (Fig. 1.2B, C). In contrast to this, amphipods show a different arrangement of blastomeres, a different sequence of cell divisions, and a clear separation of micromeres and macromeres. The primordial germ cell is not situated at the end of a cell band, but it is found in the smallest micromere of the eight-​cell stage (Figs. 1.2A and 1.5A, B). The cleavage patterns of cladocerans and copepods show some resemblance to those of euphausiaceans and dendrobranchiates. This relates to the overall appearance, the early determination of cell fates, the arrangement of blastomeres, and the cell bands (Fuchs 1914, Alwes and Scholtz 2014, Loose and Scholtz 2019) (Fig.  1.2 B, C, E, Eʹ). However, the primordial germ and mesendoderm cells are not as large as those seen in dendrobranchiates and euphausiaceans. Furthermore, the area of gastrulation, the endoderm, and the primordial germ cells are not formed at the end of a cell band, but in the region where the descendants of the two founder cells of the cell band meet (i.e., at a cross-​furrow) (see Alwes and Scholtz 2014). In contrast to this, cirriped cleavage is very different at first sight. With the exception of rhizocephalans and iblids (Scholtz et  al. 2009b), cirriped cleavage is characterized by an early ­asymmetric cleavage resulting in very unequal cell sizes (Anderson 1973, Ponomarenko 2014) (Fig. 1.2D, Dʹ). In particular, one cell is very large and contains almost all the yolk. Hence, the subsequent cleavage divisions show a set of micromeres and macromeres that resembles spiral cleavage superficially (Anderson 1973). Nevertheless, apart from the high degree of unequal blastomere size and despite some divisions, the interlocking cell bands as described in euphausiaceans and dendrobranchiates can be recognized. As in other non-malacostracans, the primordial germ cells develop in the central area of a cell band and not at the end, as in malacostracans (Ponomarenko 2014) (Fig. 1.2 D, Dʹ). Principally, partial cleavages can show a determinate pattern like holoblastic and mixed cleavages (Fig. 1.3). However, if it occurs, it is difficult to recognize because landmarks such as size, position,

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Developmental Biology and Larval Ecology and shape of the blastomere are missing. Nevertheless, there are some indications for the cladoceran Leptodora kindtii that energids show early differentiation and determination, and a specific arrangement (Samter 1900, Alwes and Scholtz 2014), but this needs further confirmation (Fig. 1.3I–​K). Mirror Images For many Crustacea with total and mixed cleavages, the phenomenon of mirror-​image eggs has been reported. This means there are two chiral types of eggs with blastomere arrangements that behave in a mirror-​image way. One can thus speak of left-​handed and right-​handed embryos. Previous descriptions are available for Cladocera, Copepoda, and, within the Malacostraca, for Decapoda, Amphipoda, and Euphausiacea [reviewed in Scholtz and Wolff (2002), Alwes and Scholtz (2004, 2014), Biffis et  al. (2009), Ponomarenko (2014), Loose and Scholtz (2019)] (Fig.  1.5E). This indicates that this may be a general phenomenon within crustaceans and is only unrecognizable in representatives with superficial cleavage, because appropriate landmarks are missing. So far, the cause and significance of mirror-​image eggs is not clear and can only be speculated. However, an effect during oogenesis is likely (e.g., the position of nurse cells or other maternal factors), which correlates putatively with the mirror symmetry of the paired ovaries. Cleavage and Body Axes The body axes of crustaceans are determined early during development. The centrifugation experiments of Kaudewitz (1950) revealed that bilateral symmetry and polarity of the body are present with beginning development of the Daphnia pulex egg. Jacobs (1925) could demonstrate this in copepods. In experiments in which one blastomere of the two-​cell stage was ablated mechanically,

Holometabola

Paraneoptera

Polyneoptera

Odonata

Ephemeroptera

Zygentoma

Archaeognatha

Diplura

Protura

Collembola

Cephalocarida

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Malacostraca

Thecostraca

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“Crustacea” Mystacocarida

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total cleavage mixed cleavage superficial cleavage

Fig. 1.6. Phylogenetic distribution of cleavage modes and reconstruction of the ground pattern of Tetraconata/​ Panarthropoda. Modified after Scholtz and Wolff (2013, Fig. 4.3).



From a Single Cell to Segmental Structures

nauplii were formed that showed more or less either the left or right body half ( Jacobs 1925). More resolution has been revealed from in vivo labeling of the four-​cell stage of various crustaceans. In all cases, there are an anterior cell and a posterior cell, and two cells giving rise to the future left and right body halves. This has been shown by in vivo cell-​labeling studies of the cirriped Austrominius modestus (Ponomarenko 2014), several dendrobranchiate decapod species (Hertzler 2015), and the amphipods Cryptorchestia garbinii (Wolff and Scholtz 2002) and Parhyale hawaiensis (Gerberding et al. 2002) (Fig. 1.5). Yet, the degree of regularity and the proportions of the clonal descendants differ between these species. Moreover, the anterior cell of Amphipoda generates only the naupliar mesoderm and the primordial germ cells, whereas it contributes to dorsal (Cirripedia) or ventral (Dendrobranchiata) ectoderm (Hertzler et al. 1994, Gerberding et al. 2002, Wolff and Scholtz 2002, Ponomarenko 2014) (Fig. 1.5). Evolutionary Relationships of Crustacean Cleavage The distribution of cleavage types according to current views of (pan)crustacean relationships suggests frequent back-​and-​forth changes between total, superficial, and mixed cleavages (see Scholtz and Wolff 2013)  (Fig. 1.6). This is even true within many crustacean subgroups such as decapods, cladocerans, and isopods (Scholtz and Wolff 2013). Furthermore, it is evident that some patterns of determinate cleavage evolved several times independently. For instance, the cleavage of amphipods seems an apomorphy for the group (Scholtz and Wolff 2002) and is very different from determinate cleavage of other malacostracans such as decapods and euphausiaceans (Scholtz and Wolff 2013). Until recently, the total determinative cleavage of various crustaceans has been compared frequently with and traced back to the determinative spiral cleavage of Annelida and Mollusca (e.g., Anderson 1973, Nielsen 2001). This view was based on the almost universally accepted Articulata hypothesis of a close relationship between Arthropoda and Annelida. The nested position of arthropods within Spiralia automatically implied a derivation of arthropod cleavage, and thus also crustacean cleavage, from the spiral cleavage mode. However, true spiral cleavage, with division spindles at alternating leiotropic (sinistral) and dexiotropic (dextral) directions, micromere quartets at the animal pole, and 4d mesoderm cells, is never realized in crustaceans (Dohle 1979, Siewing 1979, Scholtz 1997, Scholtz and Wolff 2013, Ponomarenko 2014)  (Figs. 1.2 and 1.5). All attempts to shoehorn cell division patterns and cell fates of some crustaceans into the spiral mode failed (Anderson 1973, Nielsen 2001). In contrast to this, the Ecdysozoa hypothesis postulates a close relationship between Arthropoda and Cycloneuralia. Thus, arthropods are no longer part of the Spiralia, and their cleavage no longer has to be derived conceptually from spiral cleavage, but rather is compared to the cleavage types of cycloneuralians such as Priapulida, Kinorhyncha, and Nematoda (Scholtz and Wolff 2013). In a recent reconstruction, Scholtz and Wolff (2013) arrived at the conclusion that the cleavage in the arthropod stem species followed the total mode (Fig. 1.6). However, whether this ancestral total cleavage of arthropods was determinate or indeterminate remains unclear. Scholtz and Wolff (2013) suggested that at least some aspects of a distinct cleavage pattern were present—​namely, a four-​cell stage with pairwise, somewhat shifted blastomeres resulting in cross-​furrows between two non-sister cells at both poles of the egg (Fig. 1.2). This pattern occurs in species with total and mixed cleavage modes and has been described for ostracods (Müller-​Calé 1913), anostracans (Benesch 1969), cladocerans (Kühn 1913, von Baldass 1941), cirripeds (Ponomarenko 2014), copepods (Fuchs 1914, Loose and Scholtz 2019), dendrobranchiate (Hertzler and Clark 1992) and pleocyemate (Klann and Scholtz 2014) decapods, euphausiaceans (Alwes and Scholtz 2004), anaspidaceans (Hickman 1937), and some parasitic isopods (Strömberg 1971). In many cases, this blastomere arrangement leads to two interlocking cell bands, each with a more or less parallel orientation of mitotic spindles.

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Developmental Biology and Larval Ecology In spherical eggs, this results in a pattern resembling a tennis ball (see Hertzler 2015) (Fig. 1.2). A corresponding pattern can be assumed tentatively for the ancestor of crustacean and hexapods—​the stem species of Tetraconata/​Pancrustacea (Scholtz and Wolff 2013). This renders superficial cleavage in this group as derived. Likewise, the different patterns of strict determinate cleavage evolved within crustaceans. For the latter, there is no example among the Hexapoda (Anderson 1973). Gastrulation Modes Gastrulation can be defined as the process that leads to the separation of germ layer ectoderm, endoderm, mesoderm, and the prospective germ cells. As is true for early cleavage, gastrulation of crustaceans follows a wealth of different patterns (Fioroni 1970a, Weygoldt 1979, Gerberding and Patel 2004, Wolff and Gerberding 2015). Gastrulation in crustaceans can be categorized according to the mode of cell layer formations, the position of the gastrulation center along the forming body axis, the stage at which gastrulation begins, and the various phases of germ layer formation. The mode of cell separation shows a great variation. One finds cell immigration [Cladocera: Kühn (1913), Copepoda: Fuchs (1914), Mysidacea: Manton (1928), Isopoda: Strömberg (1965)], invagination (i.e., forming a proper blastopore) [Decapoda:  Reichenbach (1886)], delamination (i.e., radial cell divisions) [Ostracoda: Müller-​Calé (1913), Decapoda: Romero-​Carvajal et al. (2018)], and epiboly (i.e., the mesendoderm cells are overgrown by ectoderm cells) [Cirripedia: Anderson (1973), Ponomarenko (2014); parasitic Copepoda: Kohler (1976)] (Fig. 1.7). Yet, the boundaries between the different internalization modes are not always very clearly set. Immigration and invagination as well as immigration and epiboly are sometimes combined [Anaspidacea: Hickman (1937); Amphipoda:  Scholtz and Wolff (2002); Decapoda:  Fioroni (1970a), Hertzler (2015)] (Fig. 1.7C, D, F, G, I, J). Likewise, the area of gastrulation differs with respect to the anterior and posterior region of the forming embryo. It has been found to be localized close to the anlagen of the mouth (stomodaeum), as in amphipods (Scholtz and Wolff 2002)  (Fig. 1.7D), or the anus (proctodaeum), as in decapods like the crayfish (Alwes and Scholtz 2006) and shrimp (Ishikawa 1885, Zilch 1978, Hertzler and Clark 1992, Hertzler 2015) (Fig. 1.7A, K). In the latter case, one can speak of a deuterostome condition that resembles the development of echinoderms and vertebrates (see Martin-​ Duran et al. 2012). The beginning of the process of gastrulation is also very variable (Fig. 1.7). The extreme examples of the stage of initial cell internalization are marked by the anaspidacean Anaspides tasmaniae, in which the first cells immigrate at the 16-​to 32-​cell stage (Hickman 1937); and the freshwater crayfish Astacus, in which cells leave the blastoderm after 1024 cells have been formed (Zehnder 1934). Regardless of cleavage mode and yolk content, most crustaceans begin gastrulation around the ­seventh cleavage (Samter 1900, Kühn 1913, Müller-​Calé 1913, Fuchs 1914, Manton 1928, Dohle 1970, 1972, Fioroni 1970a, Strömberg 1965, Scheidegger 1976, Alwes and Scholtz 2004, Browne et al. 2005, Hertzler 2015, Loose and Scholtz 2019). However, the determination of cell fates related to the various germ layers might happen much earlier. For instance, the highly unequal first cleavage division of cirripeds and parasitic copepods determine the (mes-​)endodermal fate of the large yolk-​containing blastomere (Anderson 1973, Kohler 1976, Ponomarenko 2014). Furthermore, in dendrobranchiate shrimp, in cladocerans, and copepods, natural markers such as intracellular granular material (e.g., intracellular body; see Fig. 1.7G) have been found that allow the identification of the germ line from early cleavage divisions onward (Häcker 1897, Kühn 1913, Fuchs 1914, Biffis et al. 2009, Vincent and Hertzler 2018) (Fig. 1.7C, G, I). Single-​cell labeling and blastomere isolation in amphipods has revealed that the primordial germ line, the ectoderm, the naupliar mesoderm, and the postnaupliar mesendoderm are



From a Single Cell to Segmental Structures

determined at the eight-​cell stage (Gerberding et al. 2002, Wolff and Scholtz 2002, Extavour 2005). In this case, the smallest micromere divides once more to produce the division-​arrested pair of primordial germ cells of the two body halves (Fig. 1.5A, cells al and ar). This finding has been confirmed by gene expression data (Extavour 2005, Gupta and Extavour 2013). The number of cells that form the inner germ layers varies to a great degree. After determined cleavage with a stereotyped pattern, there are often just a few identifiable mesoderm, endoderm, and germ cell precursors. Euphausiacea and dendrobranchiate Decapoda differentiate two large cleavage-​arrested cells during the formation of the 16-​to 64-​cell stage (depending on the species) that give rise to primordial germ cells, the postnaupliar mesoderm, and the endoderm. These are encircled by eight or nine crown cells that form the naupliar mesoderm (Alwes and Scholtz 2004, Hertzler 2015) (Figs. 1.2B, C and 1.7F, G). Similar numbers and a similar cell arrangement have been reported for Cladocera and Copepoda (Kühn 1913, Fuchs 1914, Alwes and Scholtz 2014, Loose and Scholtz 2019) (Figs. 1.2E, Eʹ and 1.7C). In contrast to this, numerous cells are affected by the invagination gastrula of amphipods and freshwater crayfish (Reichenbach 1886, Scholtz and Wolff 2002) (Fig. 1.7A, D, J, K). In most cases, separation of germ layers in crustaceans is a more or less continuous and coherent process involving the internalization of a mesendoderm mass, the primordial germ cells, and, in yolk-​rich eggs, the endodermal [or in cladocerans, perhaps mesodermal (see von Baldass 1941)] vitellophages or yolk cells, which are either transitory yolk-​consuming cells or are involved in midgut formation (Manton 1928, 1934, Shiino 1942, Weygoldt 1961, Strömberg 1965, Fioroni 1970b, Scheidegger 1976) (Fig. 1.7H, K). The sequence of the internalization of these germ layers can vary to some degree between crustacean groups. For instance, the gastrulation process is initiated by the formation of vitellophages in the decapod shrimp Macrobrachium carcinus and the tanaidacean Heterotanais oerstedii, whereas vitellophages form from immigrated mesendoderm cells in another shrimp, Palaemon varians, and the leptostracan Nebalia bipes (Manton 1934, Weygoldt 1961, Scholl 1963, Müller 1984). Furthermore, the mesoderm and the endoderm can immigrate independently, as is exemplified in Anostraca, some Cladocera, and Anaspidacea (Hickman 1937, von Baldass 1941, Benesch 1969). The mesendoderm mass differentiates into mesoderm and endoderm cells, but mostly it is difficult to discriminate between these two cell groups before they separate spatially (Fig. 1.7). In some cases, gastrulation is polyphasic. In other words, it involves spatial or temporal gaps between the formation of the different germ layers (Fioroni 1970a, Weygoldt 1979). For instance, in the thermosbaenacean Thermosbaena mirabilis, endodermal vitellophages immigrate into the yolk in an anterior area of the germ disk, whereas the gastrulation center sensu stricto lies in the posterior region. Here the formative mesendoderm cells immigrate that give rise to muscles, blood vessels (mesoderm), and midgut (endoderm) (Zilch 1974). A different pattern of polyphasic gastrulation has been described for the anostracan Artemia salina (Benesch 1969). In this species, the first phase of gastrulation is characterized by the immigration of mesodermal cells and primordial germ cells, the second phase follows when endoderm cells immigrate, which form the gut anlage, from a different gastrulation area (Benesch 1969). Additional cases of polyphasic gastrulation can be found among the anaspidacean Anaspides tasmaniae and several decapods (Hickman 1937, Fioroni 1970a,b, Müller 1984). The mode of gastrulation is generally thought to correlate to the yolk content of the eggs. Yolk-​ poor eggs with total cleavage and a blastula with a central blastocoel are expected to undergo an invagination gastrula. In contrast to this, yolky eggs with superficial cleavage and a yolk-​filled blastoderm show immigration or delamination gastrulation modes (Fig. 1.7). However, like the correlation between yolk content and cleavage mode, that of yolk content and gastrulation is not that straightforward. For instance, total-​cleaving ostracod eggs show a concerted delamination of a few cells (Müller-​Calé 1913), whereas the large and yolky eggs of freshwater crayfish show a clear invagination gastrula (Reichenbach 1877, 1886, Alwes and Scholtz 2006) (Fig. 1.7A, K).

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(F)

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(E)

(I) (G)

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Fig. 1.7. Gastrulation. Modes of endoderm, mesoderm, and germ cell separation. (A–​F) Superficial aspects of the gastrulation center. (G–​K) Sections through the gastrulation center. (A) The anterior end of the germ band is characterized by the head lobes (hl), posterior invagination with a blastopore (bp) close to the proctodaeum (pr) and a large endoderm (en) sack [compare with (K)]: crayfish Cherax destructor. (B) Epiboly with micromeres overgrowing (dotted line) the yolky macromere: cirriped Austrominius modestus. (C) Immigration with a slight invagination and early determination of the primordial germ cells (the arrow points to the germ granules) (green), the endoderm (yellow), and the crown cells forming mesoderm (red):  cladoceran Polyphemus pediculus. (D) Anterior invagination forming a blastopore (bp) combined with immigration [compare with ( J)]:  amphipod Cryptorchestia garbinii. (E)  Late gastrulation with immigration and formation of ectodermal teloblasts (ET) arranged in a semicircle in front of the gastrulation center (ga):  isopod Asellus communis. (F) Immigration of two large mesendoderm cells, surrounding crown cells (cc) (mesoderm) with radial mitotic spindles, and subsequent invagination forming a blastopore (bp): euphausiacean Meganyctiphanes norvegica. (G) Immigration of two large mesendoderm cells (me) [one containing the intracellular body (icm),



From a Single Cell to Segmental Structures

The Role of Cell Linage for the Differentiation of Germ Layers and Primordial Germ Cells The origin of germ cells has been traced by the expression of the vasa gene in identified cells in the amphipod Parhyale hawaiensis, in the pleocyemate decapod Macrobrachium nipponense, and in the cladoceran Daphnia magna (Extavour 2005, Sagawa et al. 2005, Özhan-​Kizil et al. 2009, Qiu et al. 2013, Gupta and Extavour 2013). Interestingly, vasa expression in amphipods begins after the primordial germ cells have been differentiated (Özhan-​Kizil et al. 2009). Thus, it is not involved in the determination of the primordial germ cells, but only in the maintenance of their fate (Özhan-​Kizil et al. 2009). Something similar appears to be the case in Daphnia magna, in which the earliest expression of vasa occurs at the eight-​cell stage. This stands in contrast to many other metazoans and the decapod Macrobrachium nipponense, in which vasa has been detected in a certain region of the zygote and in a single blastomere from early cleavage stages onward (Qiu et al. 2013). These results suggest that in cases with such highly stereotyped cell lineages as in amphipods, these lineages play a crucial role for cell fate determination (see Stent 1985). Something similar can be observed concerning the twist gene. In arthropods lacking stereotyped cell division patterns, such as spiders and insects, this gene has been shown to be expressed relatively early during gastrulation in mesoderm precursors (Price and Patel 2008). As is the case for vasa, in the amphipod Parhyale hawaiensis, twist is expressed relatively late in a subpopulation of segmental mesoderm cells (Price and Patel 2008). Hence, the determination of cell fate as mesodermal predates gene expression. The Ancestral Pancrustacean/​Tetraconate Gastrulation As a result of the great diversity, the ancestral pattern of gastrulation of Pancrustacea/​Tetraconata is almost impossible to reconstruct. If one accepts the conclusion that the ancestral cleavage mode was a total cleavage, as described earlier, the gastrulation mode might have followed a similar pattern to the one shared by some cladocerans, cirripeds, and euphausiaceans, and dendrobranchiate decapods among Malacostraca. This includes an early determination of the germ line, the immigration of a germ cell–​mesendoderm complex comprising only a few cells, and a ring-​like arrangement of crown cells. Together, these cells form a blastopore with a relatively flat invagination. However, the paraphyletic apterygote hexapods show a very different type of gastrulation, involving endoderm formation by scattered internalization of vitellophages during cleavage or from cells separating from the blastoderm. The mesoderm originates from the midline, or also from scattered internalizations from the presumptive ectoderm (see Anderson 1973). Yet, because of these differences and because data about many crustacean taxa are missing and the exact cell lineage of cladocerans and copepods still needs to be confirmed, any inference about the plesiomorphic gastrulation of Pancrustacea/ Tetraconata is necessarily preliminary. a kind of germ granule], surrounding crown cells, and, later, invagination: dendrobranchiate Penaeus monodon. (H)  Three subsequent stages (from left to right) (sagittal section). Gastrulation center (ga) with immigration forming a shallow pit. The ectodermal teloblasts (ET) differentiate at the anterior margin of the gastrulation center and produce smaller ectoderm cells in an anterior direction, primordial germ cells (gc), mesoderm (mes), and yolk containing endoderm cells (en) migrate anteriorly, mesoteloblasts (MT) bud further mesoderm cells in an anterior direction [compare with (Figs. 1.9 and 1.10)]: mysid Hemimysis lamornae. (I) Sagittal section of later stage of (C) with an almost closed blastopore (bp), showing immigration of germ cells (gc), mesoderm (mes), and endoderm (en). ( J) Cross-​section of (D) depicting invagination (bp) and cell immigration (asterisks). (K) Sagittal section of Astacus astacus; compare with (A). The endoderm (a) forms a large midgut primordium with secondary yolk pyramids, the proctodaeum (pr) elongates and forms the hindgut, and the invagination of the stomodaeum (st) begins. The large ectoteloblasts (ET) surrounding the caudal papilla and some mesoderm cells are visible. (A) Photo courtesy of Gerhard Scholtz. (B) Photo courtesy of Ekaterina Ponomarenko. (C, I) Modified after Kühn (1913, Figs. 50 and 74). (D) Photo courtesy of Gerhard Scholtz. (E) Modified after McMurrich (1895, Fig. 36). (F) Photo courtesy of Frederike Alwes. (G) Modified after Biffis et  al. (2009, Fig.  7f). (H)  Modified after Manton (1928, Fig. 3). ( J)  Modified after Scholtz and Wolff (2002, Fig. 12b). (K) Modified after Reichenbach (1896, Fig. 66). See color version of this figure in the centerfold.

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Developmental Biology and Larval Ecology Germ Disk Either during the gastrulation process or after it has been completed, superficial cells assemble to form the germ rudiment that represents the starting point for the formation of the adult body organization. As has been shown in amphipods, this process involves dramatic cell migration (Scholtz and Wolff 2002). In yolky eggs, this phase is characterized by the germ disk that marks the future ventral side of the embryo (Fig. 1.7D). The germ disk comprises outer ectodermal cells and inner layers or groups of mesodermal, endodermal, and primordial germ cells (Manton 1928, Weygoldt 1958, Browne et al. 2005). These cells are free of yolk and densely packed. In some cases, vitellophages are situated in the yolk mass, which digests the yolk (Scholtz and Wolff 2002). The remaining area of the egg surface is called the extra-​embryonic region, because it plays no formative role. In this region, the cells are large, contain yolk, and are arranged loosely. Probably some of these cells also act as vitellophages (Fig. 1.7D). With the formation of the germ disk, the anteroposterior axis of the future body becomes recognizable. The germ disk elongates and turns into the germ band. Characteristic landmarks are the anterior and posterior invaginations of the stomodaeum (ectodermal mouth and foregut) and the proctodaeum (ectodermal anus and hindgut) (Figs. 1.7K and 1.8A). At the germ band’s anterior margin, the head lobes form, which are the anlagen of the lateral eyes and some anterior brain parts (Manton 1928, Weygoldt 1958, Scholl 1963, Dohle 1972, Browne et al. 2005) (Figs. 1.7A and 1.8A). At the posterior end, the preanal growth zone is differentiated (Ooishi 1959, 1960, Dohle 1970, Scholtz 1992) (Figs. 1.8A–​1.10). This produces the ectodermal and mesodermal cellular material that elongates the germ band and eventually gives rise to the ectodermal segmental structures, such as intersegmental furrows, ganglia, and limbs, and the mesodermal musculature. During this process, the endodermal midgut precursor cells form the midgut and the midgut glands, if present (Reichenbach 1886, Fioroni 1970b, Wolff and Scholtz 2002) (Figs. 1.5B and 1.7K). With the elongation of the germ band and the differentiation of the segmental structures, the germ band extends laterally and the extra-​embryonic region gets smaller until the lateral halves of the embryo meet at the dorsal side. This dorsal closure is accompanied by the digestion of the yolk mass, which at the hatchling stage is reduced to a large degree or is gone entirely (Scholl 1963). A germ disk and the subsequent germ band are absent in species with small, almost-​yolk-​free eggs that undergo total or mixed cleavage, such as some copepods and cladocerans (Kühn 1913, Fuchs 1914). In these cases, the germ anlage comprises the entire embryo, including the dorsal side. Germ Band Shape After gastrulation and during the differentiation of the germ band, the embryos assume a characteristic shape that differs among taxa. The plesiomorphic condition within malacostracans is characterized by the formation of a transverse ventral groove: the caudal furrow (Figs. 1.8 and 1.10). With advanced development, this groove deepens and subdivides the embryo in an anterior yolky region and a posterior ventrally folded, mostly yolk-​free caudal papilla (Scholtz 1984, 2000) (Figs. 1.8A and 1.10D). The latter starts as a dome-​shaped bud in the posterior region of the embryo. In correlation with the budding of cells in the posterior growth zone, the caudal papilla elongates and becomes flexed ventrally (Figs. 1.8A and 1.10D). Because the growth zone encircles the caudal papilla, the ventral and dorsal parts of the embryo and the subsequent segments are formed. With hatching, the caudal papilla stretches backward and supports the rupture of the egg envelopes. The groove between the anterior region and the caudal papilla does not correspond to the boundary between thorax and pleon. Rather, the caudal papilla comprises the anlagen of



From a Single Cell to Segmental Structures

a (different) number of thoracic, all pleonic segments, and the telson (Scholtz 2000)  (Fig. 1.8). A caudal papilla has been found in embryos of Leptostraca, Stomatopoda, Decapoda, Syncarida, and Thermosbaenacea (Scholtz 2000). In the latter, the caudal papilla contains some yolk, but the growth zone still encircles the papilla (Zilch 1974). Among peracarids, Mysidacea have a caudal papilla, but the yolk content is relatively high and the growth of the germ band is restricted to the ventral side (Scholtz 1984). Amphipoda show a caudal groove, but the posterior body is almost as wide as the anterior part and, as in mysidaceans, only the ventral side constitutes the germ band (Weygoldt 1958, Scholtz 1990, Ito et al. 2011) (Figs. 1.8 and 1.9). In contrast to this, the embryos of Isopoda, Tanaidacea and Cumacea, Spelaeogriphacea, and Mictacea neither show a caudal groove nor a caudal papilla (Scholtz 2000, Olesen et al. 2014a) (Fig. 1.10E). The germ band grows out in one plane and only later does a dorsal furrow form, which is straightened with hatching. Accordingly, this condition has been considered an apomorphy for the Mancoidea within the Peracarida (see Richter and Scholtz 2001).

(A)

(C)

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Fig. 1.8. Egg–​nauplius and nauplius. (A) The egg–​nauplius of the crayfish Astacus astacus with the advanced anlagen of the eyes (head lobes, hl), the first and second antennae and the mandibles (a1, a2, md) compared with more posterior segmental structures. At the anterior, there is the anlage of the labrum (lr); at the posterior, there is the forming caudal papilla (cp) with the proctodaeum. (B) The nauplius larva of the northern krill Meganyctiphanes norvegica. This is a reduced nonfeeding nauplius with few cells (nuclear fluorescent stain bisbenzimide). (C) Scanning electron micrograph of an advanced embryo of the amphipod Gammarus pulex. Here an egg–​nauplius is lacking because the naupliar region shows no advanced development. The naupliar appendages are in a similar stage as the crayfish in (A); but, in contrast to this, the fifth thoracic segment shows early limb buds and the fourth pleonic segment is recognizable. cf, caudal furrow. (D) Scanning electron micrograph of the nauplius II of the cirriped Amphibalanus improvisus showing the advanced development of a proper nauplius compared with an egg–​nauplius (A) and the nauplius of the krill (B). A large labrum (lr) is present. ff, frontal filaments. (A) Modified after Reichenbach (1896, Fig. 9). (B) Photo courtesy of Gerhard Scholtz. (C) Photo courtesy of Gerhard Scholtz. (D) Modified after Semmler et al. (2009, Fig. 1b). See color version of this figure in the centerfold.

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Fig. 1.9. Camera lucida drawings of the germ bands of a cladoceran and two malacostracans. The first thoracic segment (th1) is labeled for comparison. (A) The postnaupliar germ band of the cladoceran Leptodora kindtii. There are many small cells with an irregular arrangement; ectoteloblasts are lacking. (B) Germ band of Neomysis integer. There are relatively few cells, which are arranged regularly in a grid-​like pattern of longitudinal columns (including a midline) and transverse rows. The large ectoteloblasts (arrow) give rise to regular rows that follow a determined sequence of division (yellow: undivided rows, blue: rows in the phase of the first wave of division, green: rows in the phase of the second wave of division, orange: beginning differential cleavages and morphological segmentation, red: the offspring of one ectoteloblast along the anteroposterior axis of the germ band). ie, intercalary elongation; sb, segmental border; gb, genealogical boundary (for an explanation, see the text). (C) The advanced germ band of the amphipod Gammarus pulex. In this case, the regular grid-​like pattern of ectoderm cells forms without ectoteloblasts. This is an apomorphy of Amphipoda. The midline is omitted; the mesoderm is drawn in bold lines. The eight mesoteloblasts (MT) at the end of the germ band and their transverse descendant rows are visible (numbers in brackets). Some individual cells of the mesoderm are labeled. In malacostracans the telson is formed posterior to the teloblasts. (A) Modified after Gerberding (1997, Fig. 5c). (B) Modified after Scholtz and Wolff (2013, Fig. 4.7). (C) Modified after Scholtz (1990, Fig. 36). See color version of this figure in the centerfold.

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Fig. 1.10. Teloblasts. (A) A cross-​section through the caudal papilla in the area of teloblasts of Homarus americanus. An outer circle of 19 ectoteloblasts (ET) and an inner circle of eight mesoteloblasts (MT) around the proctodaeum (pr) are visible. This is the ancestral condition within Malacostraca. (B) A similar section in the freshwater crayfish Cherax destructor. About 40 ectoteloblasts are combined with eight mesoteloblasts, both situated around the proctodaeum (pr). (C) The growth zone of the peracarid Neomysis integer. The ectoteloblasts form a transverse row of about 15 cells. (D) The germ band with ventrally folded caudal papilla of Astacus astacus. The arrow



From a Single Cell to Segmental Structures

Growth Like hexapods, myriapods, and chelicerates, non-malacostracan crustaceans do not show specific cell types in their growth zone. Furthermore, the cells in the germ band do not display a recognizable stereotyped pattern of cell division and arrangement. This is different in malacostracans. The stereotyped formation and differentiation of the malacostracan germ band (see Figs. 1.9 and 1.10) allowed very thorough studies, leading to the most detailed knowledge about the early segmentation process in crustaceans (Dohle et al. 2004). The growth zone of this group is characterized by transversely arranged, relatively large stem cells:  the teloblasts (Figs. 1.9 and 1.10). Teloblasts occur in the ectoderm (ectoteloblasts) and mesoderm (mesoteloblasts). They divide asymmetrically, producing smaller cells in an anterior direction (Figs. 1.9 and 1.10). This way, the cellular material for the subsequent segmentation of the germ band is generated. A ring of 19 ectoteloblasts (one unpaired ventral median ectoteloblast and nine paired ectoteloblasts on either side of the midline) and an inner ring of eight mesoteloblasts (four on either side; a median mesoteloblast is absent), both surrounding the caudal papilla anterior to the telson anlage, has been found in Leptostraca, Stomatopoda, most Decapoda, Euphausiacea, Anaspidacea, and Thermosbaenacea (Scholtz 2000)  (Fig. 1.10A). This distribution allows for the conclusions that this pattern is the original condition for Malacostraca (Scholtz 2000). However, within malacostracans, some evolutionary changes have taken place. In the lineage to freshwater crayfish, the number of ectoteloblasts increased to about 40, but the ring arrangement persisted (Scholtz et  al. 2009a) (Fig. 1.10B). In contrast to this, the teloblast rings have been evolutionarily transformed to transverse rows with a variable number (15–​23) of ectoteloblasts in Peracarida (Fig. 1.10C). The Amphipoda are a notable exception, because they lost ectoteloblasts entirely (Bergh 1894, Scholtz and Wolff 2002) (Fig. 1.9). In contrast to the situation in ectoteloblasts, the mesoteloblasts remain conservative and, so far, no exception to the number eight has been found. Even amphipods that lost ectoteloblasts evolutionarily are equipped with eight mesoteloblasts (Fig. 1.9). The only evolutionary change concerns the peracarids, which show a transverse row of mesoteloblasts instead of the plesiomorphic ring arrangement (Scholtz 2000). Ectoteloblasts are generated in different ways. Ooishi (1959, 1960) described a complex, stereotyped cell division pattern for a decapod shrimp, and a hermit and brachyuran crab (Fig. 1.10F). This way, the ectoteloblasts are differentiated in a stepwise manner and the most ventral ectoteloblasts begin their characteristic asymmetric divisions before the more dorsal ectoteloblasts have been differentiated. The ectoteloblasts in a freshwater crayfish and a cumacean have been reported to form in situ without any special lineage, but nevertheless in a ventral–​dorsal sequence (Dohle 1970, Scholtz 1992). In contrast, the differentiation of mesoteloblasts follows a corresponding cell lineage in decapods, cumaceans, and amphipods (Ooishi 1959, 1960, Dohle 1970, Scholtz 1990, Price and Patel 2008, Hunnekuhl and Wolff 2012) (Fig. 1.10G). With their asymmetric divisions, the teloblasts generate transverse rows or rings of descendant cells that form a regular grid-​like pattern (Hejnol et al. 2006). The divisions of the teloblasts follow more or less a mediolateral wave of mitoses on either side of the midline (Fig. 1.9). The unpaired points to the ectoteloblasts. Again, these are more than 19 in number and Reichenbach (1886) is the first record of this type of stem cells. (E) Ventral aspect of the germ band of the isopod Porcellio scaber. In this case, the germ band grows around the yolk without the formation of a caudal furrow or a caudal papilla. (F) The cell division pattern of the differentiation of ectoteloblasts in some decapod crustaceans. (G) The cell division pattern of the differentiation of mesoteloblasts in the peracarid Diastylis rathkei. A corresponding pattern has been described for amphipods and decapods. (A) Modified after Dohle et al. (2004, Fig. 3). (B) Photo courtesy of Gerhard Scholtz. (C) Photo courtesy of Gerhard Scholtz. (D) Modified after Reichenbach (1896, Fig. 10). (E) Modified after Scholtz and Wolff (2013, Fig. 4.6c). (F) Modified after Ooishi (1959, Fig. 8). (G) Modified after Dohle et al. (2004, Fig. 5a). See color version of this figure in the centerfold.

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Developmental Biology and Larval Ecology median ectoteloblast generates a midline column of unpaired cells that matches the symmetry axis of the body in the postnaupliar region (Figs. 1.9 and 1.10C). Each ectodermal transverse row or ring of ectoteloblast derivatives forms a genealogical unit (Fig. 1.9). In the ectoderm, the cells of each of these descendant rows divide twice following a mediolateral wave of mitoses with an anteroposterior spindle orientation (Dohle et al. 2004, Scholtz and Wolff 2013) (Fig. 1.9). This way, each descendant row generates four daughter rows, which are still arranged in a regular grid (Fig. 1.9). This process elongates the germ band further. Hence, germ band elongation is a two-​step process: first, the generation of ectoteloblast descendant rows (founders of the genealogical units); second, the two waves of divisions in longitudinal direction of each of these rows (Scholtz and Wolff 2013) (Fig. 1.9B, for example). After that, the differential cleavages begin. These show different spindle directions and sometimes asymmetric divisions, but these mitoses still follow a stereotyped pattern (Fig. 1.9). With these differential cleavages, the germ band becomes three-​dimensional and the segmental structures such as intersegmental furrows, limb buds, and ganglion primordia are formed (Dohle et al. 2004) (Figs. 1.9–​1.13). When the grid-​like arrangement of the ectoderm cells was detected, it was thought that each of the ectoteloblast descendant transverse rows or rings gives rise to a morphological adult segment (Bergh 1893, McMurrich 1895, Manton 1928). Yet, more detailed analyses have shown that the segmental boundaries run transversely and slightly obliquely through the progeny of each ectoteloblast descendant row (Dohle 1972, 1976). Hence, in the ectoderm, every morphological segment is formed by the progeny of two genealogical units and, likewise, segmental ganglia and legs are composite structures (Dohle et al. 2004) (Figs. 1.9, 1.11–​1.13) Interestingly enough, amphipods show the same regular pattern of the ectoderm cells in the germ band despite the absence of ectoteloblasts (Scholtz 1990). In this case, the rows form by an arrangement of previously scattered ectodermal cells in an anteroposterior sequence (Fig. 1.9). It appears that the midline cells play an organizing role during the process of row generation and segment differentiation. This is indicated by the formation of the midline propagating in an anteroposterior direction and the generation of ectoderm rows from the midline toward the lateral direction (Scholtz 1990). Experiments in which midline cells of the amphipod Parhyale hawaiensis have been ablated with laser beams, and the expression of the single-​minded (sim) gene has been suppressed, corroborated this view strongly. Both approaches led to a “dorsalization” of ventral areas of the germ band (Vargas-​Vila et al. 2010). The sim gene is known from Drosophila to play a role in the differentiation of midline and ventral cell fates (see Vargas-​Vila et al. 2010). Apparently, a mediolateral gradient of sim is also responsible for the ventrodorsal differentiation of segmental structures in the amphipod Parhyale hawaiensis (Vargas-​Vila et  al. 2010). In malacostracans with ectoteloblasts, the role of the midline must begin with the differentiation of the median ectoteloblast cell. The latter produces most of the midline cells in an anterior direction (Dohle et al. 2004). The mesoderm descendant rows show a much larger distance from each other and undergo a different sequence of mitoses. Furthermore, there is no unpaired midline population as in the ectoderm. Nevertheless, they also reveal a stereotyped arrangement and cell division pattern (Dohle 1972, Scholtz 1990, Price and Patel 2008, Hunnekuhl and Wolff 2012) (Fig. 1.9). In contrast to the ectoderm rows, each mesoteloblast descendant row generates the mesodermal equipment of one morphological segment. Hence, in the mesoderm, the genealogical units and the segments match. There is only one curious exception. It concerns the innermost paired mesoderm cells of the first thoracic segment. These are generated in the second maxillary segment and migrate posteriorly to complement the mesoteloblast descendants of the first thoracic segment to the full set of four mesoderm cells per side (Scholtz 1990, Price and Patel 2008). The segmental mesoteloblast descendant cells of each side form the ventral body musculature (innermost mesoteloblast derivatives), the muscles of the limb (the two median mesoteloblast derivatives), and the dorsal heart (the most lateral mesoteloblast derivatives) (Hunnekuhl and Wolff 2012).



From a Single Cell to Segmental Structures

The teloblasts do not generate the material for all segments. The cells of the prospective naupliar region are formed before the teloblasts are differentiated. Ectodermal and mesodermal cells seem irregularly distributed, and a stereotyped cell lineage cannot be recognized (Dohle et  al. 2004, Scholtz and Wolff 2013). In addition, the ectodermal cells giving rise to the segments of the first and second maxillae and the anterior part of the first thoracic segment do not descend from the ectoteloblasts. Nevertheless, they are arranged in transverse rows like the ectoteloblast derivatives and they show a similar further cell division pattern, with the notable exception of the anterior part of the first maxilla, which shows a slightly different cell arrangement and division pattern (Dohle et al. 2004, Wolff and Scholtz 2006). After the formation of 12 descendant rows in the cumacean Diastylis rathkei or 13 descendant rows in the decapod Cherax destructor, the ectoteloblasts quit their characteristic asymmetric divisions. Two more rows are formed with somewhat more irregular divisions, which deliver the cells for the transition between the last segment and the telson (Dohle 1970, Scholtz 1992). The postnaupliar mesoderm behaves differently. Descendants of mesoteloblast precursors and differentiated mesoteloblasts form the mesodermal equipment of the postnaupliar segments beginning with the second maxilla (Hunnekuhl and Wolff 2012). Yet, as in the ectoteloblasts, the final divisions of the mesoteloblasts are either inverted with respect to the smaller daughter cells or symmetric (Dohle 1970, Scholtz 1990). Whether derivatives of mesoteloblast contribute to the telson mesoderm is not clear. Evolution of the Stereotyped Division Pattern As mentioned earlier, and as far as is known, the regular grid-​like arrangement of postnaupliar ­ectoderm and mesoderm cells, and the elaborate stereotyped cell division pattern are restricted to malacostracan crustaceans. A closer look at some branchiopod, cirriped, and copepod species did not reveal a comparable pattern. Neither teloblasts nor the grid-​like cell arrangement has been identified (Gerberding 1997, Dohle et al. 2004, Ponomarenko 2014, Hein and Scholtz 2018 , Hein et al. 2019) (Fig. 1.9). Furthermore, the figures of Stegner and Richter (2015, Figs. 5,6) on the development of a cephalocarid species do not indicate the presences of a regular cell arrangement and teloblasts. Claims that cirripeds and anostracans differentiate teloblasts (Anderson 1967, 1969) could not be substantiated by more recent investigations (Benesch 1969, Dohle et al. 2004, Ponomarenko 2014). However, in some non-malacostracan crustaceans (Anostraca, Cladocera, Copepoda) an unpaired midline cell population has been recognized (e.g., Gerberding 1997, Dohle et al. 2004, Ungerer et al. 2012, Hein and Scholtz 2018, Hein et al. 2019). Moreover, the images of nauplii of parasitic copepods suggest that some sort of regular cell arrangement occurs (McClendon 1907). In particular, in the light of recent phylogenetic hypotheses on the sister group of Malacostraca (Regier et  al. 2010, Schwentner et al. 2017), this deserves a reinvestigation to reveal whether it shows similarities to the pattern found in malacostracans. An invariant cell division pattern in the germ band has not been found in Chelicerata, Myriapoda, and Hexapoda (Anderson 1973). Thus, it appeared somewhere within the Pancrustacea/​Tetraconata and most likely in the lineage leading to Malacostraca (Fischer et al. 2010). Furthermore, there are some indications that some aspects of this elaborated pattern evolved only within malacostracans—​ namely, in the Caridoida (Richter and Scholtz 2001). The leptostracans and stomatopods that were studied under this aspect differentiate teloblasts with their characteristic behavior (Manton 1934, Shiino 1942, Fischer et  al. 2010). Yet, they reveal more irregularities of the cell division pattern during later stages compared with those of decapods and peracarids (Fischer et al. 2010). The reasons for the evolution of the invariant cell division patterns of malacostracans are unknown. However, a possible explanation might be found in the fact that malacostracans show a relatively low number of cells in their germ bands when compared to other crustaceans and arthropods

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Developmental Biology and Larval Ecology in general. With a low number of cells, a stereotyped cell division is necessary to put the right cell in the right place at the right time to generate a specific determination of cell fate, such as a neuronal precursor or the tip of a limb bud. If there are numerous cells to form a limb or a ganglion anlage, an exact position or number of individualized cells is not required (Schnabel 1997). Hierarchy of Germ Layers There has been a long debate about whether the mesoderm determines the fate of the ectoderm or vice versa. Experimental approaches in insects revealed that the ectoderm induces segmental fates to the mesoderm cells (Bock 1942, Haget 1953). Because gastrulation in pterygote insects follows a largely different path than that of crustaceans (Anderson 1973), one could expect that different hierarchical relationships between germ layers can be found. Indeed, comparative data of malacostracans might, at first sight, speak for a leading role of the mesoderm for the differentiation and segment formation in the germ band (Scholtz 1990). Although ectoteloblasts underwent some evolutionary changes with respect to differentiation, arrangement, and number (up to total loss), mesoteloblasts behave conservatively in this respect. There is always the number of eight mesoteloblasts. Moreover, as far as is known, all mesoteloblasts are formed via a corresponding stereotyped cell division pattern (Fig. 1.10G). However, a more detailed comparison of the segmentation in the ectoderm and mesoderm layers reveals that the differentiation process of the ectoderm proceeds in advance of that in the mesoderm. Hence, the ectoderm might have an impact on the mesoderm, rather than the other way round (Scholtz 1990). This inference has been confirmed convincingly by experimental ablations of ectodermal cells that led to a distorted pattern of the mesoderm up to absence of segmental structures and gene expressions (Hannibal et al. 2012). In contrast to this, the early formation and differentiation of the mesoteloblasts and their descendant rows are independent of ectodermal influences (Hannibal et al. 2012). Segmentation In most crustacean embryos, segment formation follows a more or less anteroposterior sequence, with more anterior segments showing advanced development. In these cases, the naupliar region comprising the eye region, and the segments of the first and second antennae and of the mandibles develops first, more or less simultaneously and with a distinct developmental gap between the nauplius segments and the postnaupliar region (Fig. 1.8). This is true for species that hatch as nauplius larva, for those that hatch at a more advanced larval stage, and for most of those with direct development. In the latter two cases, one speaks of an egg–​nauplius [Scholtz (2000) but see Jirikowski et al. (2013)] (Fig. 1.8). An egg–​nauplius is not formed in amphipod, tanaidacean, cumacean, and isopod embryos. In these cases, the gradient between the naupliar and postnaupliar regions is smooth, and a distinct gap between the development of the naupliar and the postnaupliar segments cannot be observed (Scholtz 2000) (Figs. 1.8C and 1.10E). Short and Long Germ Development The stepwise addition of segments in relation to an extension of the embryonic germ band is called short germ development (Krause 1939, Scholtz 1992, Patel et al. 1994). In species with direct development or another mode of advanced hatching, the short germ mode of development can be explained by the “embryonization” of the nauplius and metanauplius stages, including the anamorphic larval development (Fig. 1.8). However, some crustacean embryos went even further by adapting a kind of long germ development (Scholtz 1992). In these cases, the germ disk is turned into an extended germ band largely based on cell rearrangement. Anteroposterior segmentation is delayed with respect



From a Single Cell to Segmental Structures (A)

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Fig. 1.11. Segment formation and segment polarity engrailed expression in Cryptorchestia garbinii. (A) Differential interference contrast image of the whole mount of a germ band showing a thoracic segment during formation. The expression of engrailed is made visible with an antibody (brown nuclei). Each transverse stripe including the midline (m) marks the posterior margin of a segment. The intersegmental furrows form posterior to the engrailed expression. The stereotyped arrangement of ectoderm cells is recognizable. (B) Drawing of the preparation of (A) with an analysis of the mitoses of the differential cleavages and the clonal composition. Lines connect sister cells. (C) Scanning electron micrograph of three segments of the embryonic thorax (th) with artificial blue staining of the cells that express engrailed. This technique demonstrates the morphogenesis of the segmental furrows and the limb buds. Lines connect sister cells. The labels mark individually identified cells. ml, midline. (A–​C) Modified after Dohle et al. (2004, Figs. 6a and 12). See color version of this figure in the centerfold.

to germ band formation and shows a very flat gradient. Hence, the segments are formed more or less simultaneously over the entire germ band. The long germ development correlates with direct development. Hence, it occurs only in species with an epimorphic developmental mode in which all segments are present at the hatching stage. Examples are cladocerans such a Daphnia (Schwartz 1973, Mittmann et al. 2014) and, to certain extent, amphipods (Scholtz 1992) (Fig. 1.8C). Based on comparative and experimental data, it has been suggested that two processes can be discriminated that are involved in germ band differentiation. One is formation and elongation of the germ band in an anterior direction. The other is the anteroposterior propagation of the subdivision of the germ band into serially repeated units (Dohle 1972, Scholtz 1992, Williams et al. 2012, Scholtz and Wolff 2013). Hence, the growth zone does not generate segments, but just the competent cellular material, which is eventually segmented. This view allows the conclusion that short and long germ development do not necessarily imply fundamentally different mechanisms, but can be the result of a heterochronic shift between germ band elongation and subsequent segment formation (Scholtz 1992, Scholtz and Wolff 2013). Segment Morphogenesis Intersegmental Furrows The first signs of forming segments are the invaginations of the intersegmental furrows, the limb buds, and the early ganglion anlagen (Figs. 1.8–​1.13). Again, the conditions in malacostracans show the most detailed resolution of these processes. Intersegmental furrows form as transverse, slightly obliquely oriented invaginations. As mentioned earlier, they appear within the descendants of each ectoderm row (Dohle et al. 2004). The findings at the cellar level of malacostracan segmentation have been corroborated at the molecular level. Namely, the expression of segment polarity genes such as engrailed (en) and wingless concurs with the morphological results on the formation of segmental boundaries. The en gene is responsible for the establishment and maintenance of a posterior segmental cell fate in Drosophila melanogaster and other arthropods such as spiders, millipedes, and crustaceans (e.g., Patel et al. 1988, Hidalgo 1998, Damen 2002, Hughes and Kaufman 2002) (Fig. 1.11). Accordingly, it is expressed in transverse stripes in the posterior region of forming

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Developmental Biology and Larval Ecology segments. This has also been shown for a number of crustacean representatives such as decapods, cirripeds, isopods, amphipods, mysids, ostracods, copepods, and branchiopods (e.g., Patel et  al. 1988, Manzanares et al. 1996, Scholtz and Dohle 1996, Abzhanov and Kaufman 2004, Deutsch et al. 2004, Wolff 2009, Ikuta 2018, Hein et al. 2019). In all these species, en is expressed at the posterior margin of forming segments (Fig. 1.11). Yet, knowledge of the cell lineage in the germ band of malacostracans allows for an unmatched cellular resolution of en expression. It has been shown that en is expressed in the anteriormost cells of the ectodermal genealogical units after the second round of mediolateral divisions just in front of the forming intersegmental furrows (Scholtz and Dohle 1996) (Fig. 1.11). Hence, the results of lineage analyses that the anterior cells of a genealogical unit contribute to the posterior region of a morphological segment have been corroborated by the en marker. Moreover, this result shows that the genealogical units of malacostracans can be compared to the insect parasegments—​in other words, fundamental initial developmental units that subdivide the anteroposterior body axis into repeated structures that are offset to the morphological segments (Lawrence 1992). Limb Formation Two descendant rows anterior to the intersegmental furrow and in the third cell from the midline, the first cells occupy a slightly elevated position and begin the formation of the limb bud (Dohle 1976, Hejnol and Scholtz 2004, Wolff and Scholtz 2008, Wolff et al. 2018) (Figs. 1.9B, C and 1.12A). This is the case for uniramous thoracopods and for biramous mouthparts and pleopods. Only later does the limb bud of the biramous appendages widen and form the two tips of the endo-​and exopod (Fig. 1.12A, B). Like other segmental structures, the ectoderm of the limbs is composed of the cells of two genealogical units (Dohle 1976, Hejnol and Scholtz 2004, Wolff and Scholtz 2008, Wolff et al. 2018) (Fig. 1.12A–​C). A clonal analysis of the legs of the amphipod Cryptorchestia garbinii reveals that the clones of the two branches of the pleopods are all found in the uniramous thoracopods in a similar topological relationship stretching along the proximodistal axis (Wolff and Scholtz 2008) (Fig. 1.12C). Furthermore, the coxal plate and the epipodite are formed mainly by basal clones (Fig. 1.12C). Similar results were reported for the thoracopods of another amphipod species, Parhyale hawaiensis (Wolff et al. 2018). This distribution pattern leads to some well-​founded conclusions: (1) the one branch of uniramous limbs, and the exopod and endopod of biramous limbs share the same proximodistal axis; (2) epipodites are lateral outgrowths with separate axes; and (3) in ontogenetic and perhaps evolutionary terms, the uniramous limb might be result of a suppressed subdivision into two branches, rather than the loss of the outer branch of the exopodite. Fig. 1.12. Early limb formation. (A) Clonal composition of the early biramous limb bud of the left second maxilla of the cumacean Diastylis rathkei. Again, this morphogenesis correlates with the differential cleavages of the malacostracan germ band. The arrow marks the genealogical boundary between the descendants of two ectoderm rows. (B) Scheme of the expression of distal-​less in a thoracic uniramous and a pleonic biramous limb of Cryptorchestia garbinii. The empty circles indicate the loss of a previous expression. This exemplifies the similar anlage of uniramous and biramous limb buds and the later differentiation resulting from a loss of distal-​less expression. (C) Clonal analysis of uniramous thoracic and biramous pleonic limbs of the same species using the fluorescent vital marker DiI. All clones found in in the biramous limb along the long axis from the protopod (prp) to the tips of the exo-​and endopods (exo, endo) are distributed similarly in the uniramous limb (cx, coxa; ba, basis; is, ischium; me, merus; ca, carpus; pro, propodus; da, dactylus), suggesting that uniramous limbs are the result of the suppression of the bifurcation of the limb buds. The exo-​and endopods of limbs are formed by the same proximodistal axis. Exites such as gills (gi) and coxal plates (cxp) are the result of additional axis formations. The cells adjacent to the midline (light and dark gray 1,2) form part of the sternite (ster); those at the laterodorsal margin of the germ band (blue and lilac 8,9) contribute to the tergite (ter). (D) Embryo of the branchiopod Cyclestheria hislopi with distal-​less expression in the lobes of the limb anlagen, the labrum, and the furca. (A, B) Modified after Dohle et al. (2004, Figs. 10 and 14). (C) Modified after Wolff and Scholtz (2008, Figs. 3e and 4e). (D) Photo courtesy of Gerhard Scholtz. See color version of this figure in the centerfold.

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Developmental Biology and Larval Ecology The second and third mesoteloblast descendants of each row form the limb musculature (Scholtz 1990, Hunnekuhl and Wolff 2012). With the outpouching of the limb bud, the cells migrate in the leg anlage. Comparable studies on other crustacean taxa do not exist. The limbs of copepods and remipedes form in an overall similar way, but little is known about their cellular composition (Koenemann et al. 2009, Hein et al. 2019). The limb buds of branchiopods are elongated mediolaterally and occupy most of the segmental primordia (Fig. 1.12D). Along this extended area, the various lobes of the phyllopodous limb forms, and eventually it is raised from the body by lateral growth (Olesen et al. 2001, Pabst and Scholtz 2009, Fritsch et al. 2013). Interestingly enough, it has been shown that the stenopodous limbs of the raptorial cladoceran Leptodora kindtii show the same initial bud as the various lobes of other branchiopod limbs. Yet, instead of forming a leaf-​like limb, the lobes turn into the podomeres of an articulated leg comparable to those of eumalacostracans (Olesen et al. 2001). These morphological results on limb bud formation have been corroborated at the level of gene expression data. The distal-​less gene (dll) plays a crucial role for the formation of limb buds and the maintenance of the distal fate of forming limbs. In addition, it is found in the formation of sensory and nerve cells (Mittmann and Scholtz 2011). Besides other arthropod groups, it has been studied in a number of crustaceans as diverse as anostracans, cladoceromorphans, decapods, amphipods, isopods, and leptostracans (e.g., Panganiban et  al. 1995, Scholtz et  al. 1998, 2009b, Williams 1998, Olesen et al. 2001, Hejnol and Scholtz 2004, Pabst and Scholtz, 2009, Ito et al. 2011). Hence, it is found in the cells that initiate limb bud formation and in the forming tip of the growing limb (Fig. 1.12B, D). Again, the most detailed resolution at the cellular level has been shown for malacostracans. The dll gene product is expressed initially in the cell that also initiates limb bud growth morphologically (Hejnol and Scholtz 2004). This is true for the anlage of uniramous thoracopods and biramous pleopods. Later, more cells express dll and the biramous limb anlagen widens and forms two distal tips, which are both dll positive (Fig. 1.12B). These data complement those on cell lineage analysis of the respective limbs (Wolff and Scholtz 2008, Wolff et al. 2018). In branchiopods dll is expressed initially in the endopod and exopod, followed by all forming inner lobes of the elongated limb buds regardless of whether the adult limb is of phyllopodous or stenopodous type (Williams 1998, Olesen et al. 2001) (Fig. 1.12D). The only exception are the epipodites, which do not express dll (Williams 2004). In this case, the expression is probably more related to the forming sensory setae rather than indicating the outgrowth of the endite lobes. Ganglion Anlagen The formation of the segmental ganglia of the trunk of a number of crustaceans begins with the differentiation of a specific type of neural precursor cells: the neuroblasts (Fig. 1.13). Neuroblasts are relatively large stem cells that undergo several cycles of asymmetric divisions in one direction (Scholtz 1992). This way, they bud off smaller daughter cells into the interior of the embryo (Fig. 1.13). The daughter cells are called ganglion mother cells and they give rise to neurons and glia by a symmetric division. This mode of neurogenesis has been described for a number of crustacean and hexapod taxa, and it is considered an apomorphy of the Tetraconata. Among crustaceans, aspects of this mode of neurogenesis have been shown to different degrees in malacostracans, branchiopods, copepods, and probably cephalocarids (Bergh 1893, McMurrich 1895, Schimkewitsch 1896, Dohle 1976, Gerberding 1997, Scholtz 1990, 1992, Duman-​Scheel and Patel 1999, Harzsch 2001, Dohle et al. 2004, Müller 2006, Ungerer and Scholtz 2008, Ungerer et al. 2011, Stegner and Richter 2015, Hein and Scholtz 2018) (Fig. 1.13). All these groups show neuroblasts with more or less asymmetric divisions. However, the pattern of neuroblast differentiation, a map of neuroblasts, and the fate of ganglion mother cells is only known in more detail from some malacostracan species (Dohle et al. 2004, Ungerer and Scholtz 2008). In the amphipod Cryptorchestia garbinii, even the lineage of identified neuroblasts

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Fig. 1.13. Early neurogenesis. (A) Sagittal section through the nauplius of the copepod Tigriopus californicus. Anterior to the left. The forming segmental ganglia (dotted lines) are depicted and the first longitudinal fiber tracts (connectives) (arrowhead) are visible. At the ventral side, neuroblasts (asterisks) are formed. Neural stem cells that produce offspring (ganglion mother cells) are arranged as columns into the interior of the larva. In the end,

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Developmental Biology and Larval Ecology Fig. 1.13. continued. the nerve cells (dots) differentiate. (B) Ventral aspect of the anlage of a segment of the amphipod Gammarus pulex during differential cleavages that lead to the differentiation of superficial neuroblasts, which in turn give rise to ganglion mother cells. In the left body, half the superficial cells are omitted and only ganglion mother cells are visible. The median neuroblast is also omitted. (C–​E) Neuroblasts in the amphipod Cryptorchestia garbinii. (C) The neuroblast map (ventral view) with all identified lateral neuroblasts. The median neuroblast is omitted. (D, E) The clone of an identified neuroblast (b1hn) that generates the pioneer neurons pCC and aCC, known from Drosophila. (D)  Schematic drawing. (E)  The preparation with DiI.  ac, anterior commissure; pc, posterior of a ganglion. Other labels indicate identified individual cells after differential cleavages as in Fig. 1.11. (F, G) Neuroblasts in the branchiopod Triops cancriformis. (F) Transverse section through a larval trunk segment. Ventral side at the bottom, with lateral neuroblast (arrows) and a median neuroblast. On top of each neuroblast there are columns of smaller ganglion mother cells. (G) Scheme of neuroblasts (asterisks) and the columns of smaller ganglion mother cells. Gray: mesoderm. (A) Modified after Hein and Scholtz (2018, Fig. 2b). (B) Modified after Dohle et al. (2004, Fig. 9). (C–​E) Modified after Ungerer and Scholtz (2008, Figs. 3 and 5). (F, G) Modified after Müller (2006, Figs. 12a and 15). See color version of this figure in the centerfold.

to identified pioneer neurons (see Whitington 2004) has been traced (Gerberding and Scholtz 1999, 2001, Ungerer and Scholtz 2008) (Fig. 1.13C–​E). It has been shown that there are median and lateral homologous neuroblast–​neuron lineages in crustaceans and hexapods that lead to a homologous set of pioneer neurons (Gerberding and Scholtz 1999, 2001, Ungerer and Scholtz 2008) (Fig. 1.13C–​E). Neuroblasts of malacostracans are differentiated with the differential cleavages in the germ band (Fig. 1.13B). They do not appear simultaneously, but in a certain species-​specific sequence. In contrast to hexapods, in which neuroblasts delaminate from the ectoderm before they give rise to ganglion mother cells, crustacean neuroblasts remain in the ectoderm layer throughout their activity. With the advanced generation of neurons, the axon scaffold of the ladder-​like central nervous system comprising longitudinal connectives and transverse commissures in the neuromeres is established (Fig. 1.13A, E).

CONCLUSIONS AND PERSPECTIVES It is obvious that our knowledge about crustacean embryology is patchy. Few groups are relatively well studied under various aspects, but from others we still lack any data. In some cases, this is understandable, because the species are difficult to get or to maintain in the lab. On the other hand, it is astonishing that there are no proper analyses of the early cleavage of branchiopod Notostraca, Laevicaudata, and Spinicaudata, even though these animals are widespread and easy to handle. Against the background of the recent phylogenetic hypotheses about the internal relationships of Tetraconata/​Pancrustacea, increased knowledge of the ontogeny of particular groups is of great interest. One of the hotspots concerns the transition between crustaceans and hexapods. A study of the embryology of Branchiopoda, Remipedia, and Cephalocarida has the potential to shed more light on hexapod origins. Additional investigations on the embryology of Thecostraca and Copepoda may help to understand the morphological evolution of Malacostraca. In addition, a more comprehensive knowledge of the early ontogeny of Mystacocarida and Ostracoda would allow the reconstruction of the ancient developmental pathways of Tetraconata/​Pancrustacea as a whole.

ACKNOWLEDGMENTS I am grateful to the members of my group Vergleichende Zoologie (Comparative Zoology) for many discussions. I thank the cluster of excellence “Image Knowledge Gestaltung” project: Dynamic Form and the Einstein Stiftung, Berlin for funding.



From a Single Cell to Segmental Structures

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2 CRUSTACEAN LIMB MORPHOGENESIS DURING NORMAL DEVELOPMENT AND REGENERATION

Anastasios Pavlopoulos and Carsten Wolff

Abstract Crustaceans have been favored in developmental biology for the study of the diversification of body plans and their associated appendages, which exhibit remarkable diversity within and between species. Until recently, because of technical limitations, crustacean studies were restricted in scope to the comparison of appendage morphologies and expression patterns of candidate limb patterning genes already known from classic developmental animal models. To remedy this limitation and explore their full potential, a few select crustacean experimental models have been reinforced with powerful genomic and transcriptomic resources, new methods for forward and reverse genetic investigations, and for live imaging of entire embryos, or cell and tissue-​specific markers, with exceptional spatial and temporal resolution. These models include the malacostracan amphipod Parhyale hawaiensis and the branchiopod cladocerans Daphnia magna and Daphnia pulex, which display collectively all the different uniramous, biramous, and phyllopodous crustacean limb types. Within the past couple years, important discoveries have been made on the molecular and cellular basis of embryonic limb development and postembryonic limb regeneration. In Parhyale alone, gain and loss-​of-​function studies of Hox genes have revealed the combinatorial logic used by these genes for appendage specialization, whereas the reconstruction of single-​cell-​resolution fate maps of developing and regenerating appendages have identified the lineage restrictions and cellular behaviors driving both morphogenetic processes. Century-​old questions regarding the conservation and divergence of appendage patterning mechanisms across arthropods and bilaterians, or how these mechanisms can be used and reused throughout the lifetime of an organism, can now be addressed productively with crustaceans.

Developmental Biology and Larval Ecology. Edited by Klaus Anger, Steffen Harzsch, and Martin Thiel. © 2020 Oxford University Press. Published 2020 by Oxford University Press.

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INTRODUCTION A major driving force for the success of arthropods colonizing Earth has been their ability to diversify their appendages and adapt to immensely variable aquatic, terrestrial, and aerial environments. Among extant arthropods, crustaceans display the biggest diversity in body plans and associated appendages that have evolved striking specializations in form and function, observed both between species and within single species. Thus, crustaceans offer exceptional raw material to study the molecular and cellular basis of limb development, diversification, and, in many cases, regeneration. Until recently, developmental studies in crustaceans were restricted to morphological descriptions and expression analyses of candidate genes known from studies in the prime insect genetic model Drosophila melanogaster. In a few instances, researchers have been able to test their hypotheses functionally by posttranscriptional silencing of targeted genes by RNA interference (RNAi) (Sagi et al. 2013). Among the various crustacean models that have been used to study appendage development, a few species have attracted extra attention thanks to their biological and technical qualities:  the marine malacostracan amphipod Parhyale hawaiensis (Stamataki and Pavlopoulos 2016) and the freshwater branchiopod cladocerans Daphnia magna and Daphnia pulex (collectively referred to as Daphnia hereafter) (Ebert 2011). These crustaceans have proved to be genetically tractable and amenable to an ever-​expanding experimental repertoire of increasing scope and sophistication beyond RNAi. They are also optically tractable model organisms that have enabled, in the case of Parhyale, to capture the cellular events contributing to appendage development and regeneration in live-imaged crustaceans (Alwes et al. 2016, Wolff et al. 2018). Together with the availability of well-​assembled and annotated genomes for Parhyale and Daphnia, these recent breakthroughs in crustacean developmental biology have started offering novel insights into the decades-​old problem of the conservation and divergence of limb patterning mechanisms in arthropod lineages and across the metazoans. This chapter first provides an overview of the morphologies of the different crustacean appendage types, and summarizes some key recent advancements in developmental genetic and cell biology approaches taken in crustacean models, with an emphasis on Parhyale and Daphnia. It then covers the genetic basis of pancrustacean appendage development, focusing primarily on limbs in the trunk during the early specification of the limb primordia, the patterning and segmentation of the main limb proximodistal (PD) axis, and the diversification of limb morphology along the anteroposterior (AP) axis between crustacean lineages and within single species. Finally, it gives an account of recent discoveries made about the cellular basis of limb morphogenesis in Parhyale during normal embryogenesis and postembryonic regeneration. Besides presenting the current state of our knowledge (and ignorance) of the molecular and cellular mechanisms operating in direct-​ developing pancrustacean and arthropod limbs, and how these compare to Drosophila indirect-​ developing limbs, this chapter also offers a number of testable hypotheses that we hope will inspire and invigorate future research in the field.

CRUSTACEAN APPENDAGE MORPHOLOGY Generalized Appendage Types in Crustaceans The jointed crustacean appendages have diversified in form to serve many functions, including sensation, feeding, locomotion, reproduction, respiration, and defense. All the variations observed in extant crustaceans are believed to have originated from euarthropod ancestors with a series of similar postantennular biramous limbs composed of a proximal part bearing an inner (ventral) branch



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Fig. 2.1. Crustacean appendage types. Schematic representations of (A) a hypothetical ancestral appendage, (B) generalized biramous limb with primary and secondary branches, (C) malacostracan biramous pleopod, (D) malacostracan uniramous pereonic/​thoracic limb, and (E)  branchiopod phyllopodous limb. prp (in gray), protopod; endo, endopod; exo, exopod; telo, telopod; en, endite; ex, exite; cx, coxa; ba, basis; is, ischium; me, merus; ca, carpus; pro, propodus; da, dactylus.

and an outer (dorsal) branch (Fig. 2.1A) (Boxshall 2004, Haug et al. 2013, Jockusch 2017). This ancestral appendage type gave rise to the three main limb forms observed in extant crustaceans. Biramous Limb Type The most common type is the biramous limb (Fig. 2.1B). It consists of a proximal element called the protopod or protopodite, which generally comprises two segments: the proximal coxa and the distal basis. Some authors have also argued for the existence of a third proximal-​most segment: the precoxa (Fig. 2.1B, dotted line) (Boxshall 2004). There is still disagreement on whether this structure represents an extension of the body wall, a subdivision of the coxa, or a true limb joint. The protopod can develop a variable number of unsegmented inner (ventral) or outer (dorsal) lobes called endites and exites, respectively (Fig. 2.1B). Exites that have a (pre)coxal origin are also called epipodites. In biramous limbs, two branches extend distally from the protopod:  an inner endopod or endopodite (a.k.a. telopodite) and an outer exopod or exopodite. The endopod is typically multisegmented with up to six primary segments (subdivisions with intrinsic muscle attachments). The exopod represents the second segmented branch dorsal to the endopod. It normally has one or two primary segments but can have terminal flagella with numerous annulations (subdivisions lacking intrinsic muscles). Aquatic malacostracan crustaceans, like Parhyale, often possess biramous swimming limbs with similar endopodal and exopodal branches (Fig. 2.1C).

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Developmental Biology and Larval Ecology Uniramous/​Stenopodous Limb Type The exopod is often missing from some limbs in certain crustacean lineages (Schram 1986, Boxshall 2004)  resulting in a uniramous (a.k.a. stenopodous) limb type, like in most malacostracans, ostracods, and some raptorial cladoceran branchiopods. Uniramous limbs retain the proximal protopod with the coxa and basis, and the distal telopod (endopod) comprised of five limb segments:  named (from proximal to distal) ischium, merus, carpus, propodus, and dactylus (Fig. 2.1D). The protopodal segments usually possess endites and exites, such as protective coxal plates, gills, or special structures for brood care (oostegites). An ontogenetic stepwise reduction of the exopodal branch takes place during the indirect larval development of many decapod crustaceans as the thoracic appendages metamorphose from swimming to walking structures during lifestyle transitions. In isopods, all early thoracic limbs display an extra ​small protrusion laterally (interpreted as a vestigial exopod) that eventually integrates into the basis of the protopod (Wolff 2009). Amphipod crustaceans, like Parhyale and Orchestia cavimana, exhibit uniramous stenopodous limbs in their pereon/​thorax, and biramous limbs in their pleon/​abdomen (Ungerer and Wolff 2005, Browne et al. 2005, Wolff and Scholtz 2008). Polyramous/​Phyllopodous Limb Type The third type is the polyramous or phyllopodous limb found primarily in Branchiopoda but also in Leptostraca (Malacostraca). The phyllopodous limb is a flattened leaf-​like appendage in which the limb branches (endopod, exopod, endites, exites) exhibit a radial arrangement (Fig. 2.1E). These paddle-​like limbs are equipped with setae along their edges and are usually used for both swimming and feeding. The endopod and exopod develop as flat lobes with a low, if any, degree of segmentation (Schram 1986). Few branchiopod groups appear to have a segmented endopod, but it is not clear whether these represent primary segments or annulations (Pabst and Scholtz 2009). This secondarily flattened appearance makes it challenging to compare phyllopodous limbs with the other crustacean and arthropod limb types. General Aspects of Appendage Development in Crustaceans A common feature of crustacean development is the so-​called germ band, a thickened area of aggregated blastoderm cells corresponding to the embryo proper (Scholtz and Wolff 2013). After gastrulation (formation of the germ layers), the germ band extends longitudinally, marking the prospective ventral side of the embryo, and becomes progressively subdivided into distinct body segments (Fig. 2.2A, H). Segment and appendage formation proceeds from anterior to posterior; as the embryo (or nauplius larva) adds new segments at the posterior end, the previously formed, more-​anterior segments and appendages are developmentally more advanced than the posterior ones. Appendage buds bulge out ventrally in the same anterior-​to-​posterior progression (Fig. 2.2A–​ E, I–​L). This process requires the remodeling of the flat ectodermal epithelium of the limb primordium into the three-​dimensional bulge of the outgrowing limb (discussed later). Limb outgrowth differs between stenopodous and phyllopodous limbs. In stenopodous limbs, the original buds elongate along their primary PD axis and become progressively subdivided into an increasing number of elements until they attain their final number of limb segments (Fig. 2.2B–​ F). During the elongation phase, secondary limb branches, the endites and exites, bulge out and differentiate on the protopod (Fig.  2.2F, G). In the case of phyllopodous limbs, the early bud is shorter and broader, occupying almost the entire mediolateral width of the hemisegment (Fig.  2.2I). The first visible indentation represents the split between the exopod and endopod (Fig.  2.2J), followed by additional medial notches creating the ventral endites (Fig.  2.2K). The

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Fig. 2.2. Appendage development in Parhyale hawaiensis and Daphnia magna. (A–​G) Reconstruction of Parhyale embryogenesis (lateral views) and (H–​N) Daphnia embryogenesis (ventral views) from light-​sheet fluorescence microscopy recordings of transgenic embryos staged according to (Browne et al. 2005, Mittmann et al. 2014). Schematic representations of developing Parhyale and Daphnia thoracic limbs are shown at each stage, with the arrows indicating their growing proximodistal axis. An1, first antenna; An2, second antenna; Mn, mandible; Mx1, maxilla 1; Mx2, maxilla 2; T1–​T8, thoracic appendages; P1–​P6, pleonic/​abdominal appendages 1 to 6; Sto, stomodeum; A, abdomen; prp, protopod; endo, endopod; exo, exopod; telo, telopod. The arrowheads and asterisks indicate the exites (coxal plates and gills, respectively) in Parhyale T2 through T4 limbs. The small arrow, arrowheads, and asterisks indicate the exopod/​endopod split; the notches between endites, and the exite/​gill, respectively, in Daphnia T2 limb. The ventral midline is denoted with dotted lines.

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Developmental Biology and Larval Ecology elongating limb first extends toward the ventral midline and then folds out like a blade out of a pocketknife (Fig. 2.2L–​N). Cell proliferation and cell shape changes have been implicated in phyllopodous limb outgrowth (Freeman 2005). Experimental Models and Approaches to Study Crustacean Appendage Development Parhyale and Daphnia have stood out as powerful model systems to study the three types of ­crustacean limbs (Fig. 2.3). Both are direct developers with their body parts forming during the 10  days or 3  days of their embryogenesis, respectively (Fig.  2.2), when several experimental manipulations are possible. From its original introduction in the lab during the late 1990s, Parhyale was meant to tackle fundamental questions in developmental biology (Stamataki and Pavlopoulos 2016). Each Parhyale embryo develops 19 pairs of appendages along the AP axis that display at least 11 distinct morphologies differing in size, shape, and pattern (Fig.  2.3A) (Browne et  al. 2005, Serano et  al. 2016). Embryogenesis and early cell lineages have been described in detail for Parhyale and the other amphipod Orchestia (Gerberding et  al. 2002, Wolff and Scholtz 2002, Browne et  al. 2005, Alwes et al. 2011). Parhyale embryos are amenable to various experimental techniques and resources that are continuously generated at a fast pace. Experimental manipulations include cell microinjection, isolation, and ablation (Extavour 2005, Price et  al. 2010, Kontarakis and Pavlopoulos 2014); cell lineage tracing (Gerberding et al. 2002, Alwes et al. 2011, Konstantinides and Averof 2014, Wolff et al. 2018); in situ hybridization and immunohistochemistry (Rehm et al. 2009a,b); gene knockdown by RNAi or morpholinos (Liubicich et al. 2009); transient and stable genetic transformation with transposon and integrase-​based vectors (Pavlopoulos and Averof 2005, Kontarakis et al. 2011b, Kontarakis and Pavlopoulos 2014); conditional gene misexpression for gain-​of-​function studies (Pavlopoulos et al. 2009); gene trapping for unbiased genetic screens and targeted modification of trapped loci (Kontarakis et al. 2011b, Kontarakis et al. 2011a); and targeted genome editing with the

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Fig. 2.3. The Parhyale hawaiensis and Daphnia magna body plans. Adult (A)  Parhyale and (B)  Daphnia imaged with microcomputed tomography. An1, first antenna; An2, second antenna; T, pereonic/​thoracic appendages; P, pleonic/​abdominal appendages. The drawings of Daphnia appendages were reproduced from Cannon (1932).



Crustacean Limb Morphogenesis

clustered regularly interspaced short palindromic repeats (CRISPR)/​CRISPR-​associated (Cas) system for gene knock-out and knock-​in approaches (Kao et al. 2016, Serano et al. 2016, Martin et al. 2016). Furthermore, Parhyale has the right optical properties for live microscopic inspections of embryonic limb development and postembryonic limb regeneration over several days with exceptional spatiotemporal resolution (Alwes et  al. 2016, Wolff et  al. 2018). The huge Parhyale genome, estimated at 3.6 gigabases, has been assembled efficiently with a combination of Illumina and Dovetail-​based strategies (Kao et al. 2016). Together with the assembled transcriptomes and other genome-​wide resources (Parchem et al. 2010, Kao et al. 2016), these tools have accelerated the functional characterization of coding and noncoding sequences in Parhyale. The water flea Daphnia is an older model organism used in ecological, environmental, and evolutionary research that is becoming increasingly attractive for developmental studies (Ebert 2011). Daphnia develops two pairs of antennae (the first uniramous and the second biramous) and three pairs of gnathal appendages in the head, five pairs of phyllopodous limbs with variable morphologies in the thorax, and no limbs in the abdomen (Fig. 2.3B). Daphnia reproduce primarily asexually, making it ideal for cost-​effective maintenance of clonal populations. Detailed staging systems have been introduced for embryonic development (Mittmann et al. 2014, Toyota et al. 2016), and many experimental resources have been established for developmental genetic research. These include protocols for gene expression analyses (Shiga et al. 2002), gene knockdown by RNAi (Kato et al. 2011, Hiruta et al. 2013), transient and stable genetic transformation (Kato et al. 2012, Nakanishi et al. 2016, Nong et al. 2017), targeted genome editing with the CRISPR/​Cas system or transcription activator-​like effector nucleases (TALENs) (Hiruta et al. 2014, Nakanishi et al. 2014, Naitou et al. 2015, Kumagai et al. 2017, Hiruta et al. 2018), and live imaging (Fig. 2.2H–​N) (Nong et al. 2017). Daphniids were the first crustaceans to have a genome sequenced with an estimated size of 200 megabases. Several more resources have been generated for environmental genomic and population genetic studies, including RNA-​Seq data sets, microarray platforms, and genetic maps (Orsini et al. 2016, Dukic et al. 2016). Besides Daphnia and Parhyale, we also refer to other crustacean species used in developmental studies to understand the organization and evolution of arthropod body plans and associated appendages, primarily at a descriptive level. These species are representative of all major crustacean lineages, including branchiopods (anostracans and notostracans), malacostracans (leptostracans, mysids, isopods, amphipods, and decapods), thecostracans (cirripedes), copepods (cyclopoids), and oligostracans (mystacocarids).

GENETIC BASIS OF APPENDAGE DEVELOPMENT Appendage Specification and Early Subdivision Drosophila has served as the prime model to understand arthropod appendage specification, patterning, growth, and differentiation (Morata 2001, Kojima 2004, Estella et al. 2012). These processes take place at different stages of the Drosophila life cycle (Fig. 2.4). Limb primordia are specified as small clusters of cells during early embryogenesis, shortly after the metameric subdivision of the embryo (Fig. 2.4A) (Cohen 1990). Each thoracic cluster separates into ventral (leg) and dorsal (wing or haltere) primordia that form subepidermal sac-​like structures in the larva, called imaginal disks (Fig. 2.4B) (Cohen et al. 1993). Imaginal disks grow by cell proliferation and undergo extensive patterning during larval stages establishing the adult limb and body wall fates (Fig. 2.4C–​E). After pupation, the disks turn inside out (disk eversion) so that limbs are positioned outside the adult body wall (von Kalm et al. 1995). Differentiation of the actual adult limb structures and cuticle deposition take place during pupa metamorphosis (Fig. 2.4F).

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Fig. 2.4. Genetic basis of Drosophila thoracic limb development. Schematic representations of (A)  early specification and (B) subdivision of the embryonic limb primordium, (C) proximodistal patterning, (D) dorsoventral patterning and (E) leg gap gene expression in the leg imaginal disk, and corresponding subdivisions in (F) the adult segmented leg. Please refer to the text for gene abbreviations. Positive and negative regulatory interactions are indicated with arrows and T-​bars, respectively. Stages A and B are likely Drosophila innovations and are not applicable to crustacean limb development. The patterning mechanisms for early limb specification and subdivision in crustaceans are not clear yet. Leg gap genes together with epidermal growth factor receptor and Notch signaling, and their downstream target genes likely have conserved roles in limb proximodistal patterning and segmentation across the arthropods.



Crustacean Limb Morphogenesis

In most other pancrustacean and arthropod groups, appendages are direct three-​dimensional outgrowths of the embryonic (or postembryonic) body wall similar to the vertebrates. The embryo hatches with functional differentiated limbs that are either miniature versions of the adult limbs or undergo varying degrees of modification before they assume their final morphology. Despite this caveat, comparative studies have demonstrated that the genes involved in appendage development are similar between Drosophila, crustaceans, and other studied arthropods (Angelini and Kaufman 2005b). In this section, our knowledge of Drosophila limb development is the starting point, followed by the relevant information from crustacean and other arthropod lineages. There is a hot debate about the origin of the dorsal wings in insects and whether they have evolved from the proximal branch of an ancestral crustacean leg (like a lobe-​shaped epipodite) or have emerged as novel lateral outgrowths of the dorsal body wall (tergum) or have a dual origin (Clark-​Hachtel and Tomoyasu 2016). We concentrate on the ventral Drosophila appendages that are clearly homologous to crustacean appendages. The Limb Patterning Gene Distal-​less The gene Distal-​less (Dll; the Drosophila genetic nomenclature is followed throughout, with gene names/​abbreviations and transcribed RNAs written in italics and translated proteins written in plain type) encodes a homeodomain transcription factor with a conserved role in appendage development (Panganiban et al. 1997). It is first expressed in the limb primordia and later in the distal parts of developing limbs in all arthropods (Panganiban et al. 1995). Depending on the severity of Dll mutations, adult Drosophila develop truncated legs retaining the proximal protopod, but with reduced and fused or completely eliminated elements of the distal telopod (Panganiban 2000). Similar phenotypes have been described in crustaceans after genetic perturbations of Dll. Shortened appendages have been obtained after Dll knockdown by RNAi in stenopodous and phyllopodous appendages of Daphnia (Kato et al. 2011, Hiruta et al. 2013), and in uniramous and biramous appendages of Parhyale (Liubicich et  al. 2009). More severe phenotypes have been obtained after Dll knockout with CRISPR/​Cas or TALENs in these species, including elimination of the entire telopod (Hiruta et al. 2014, Kao et al. 2016, Hiruta et al. 2018). Besides its key role in limb specification and PD patterning, Dll has also another conserved, presumably ancestral, role in the development of the peripheral sensory organs in proximal and distal positions of arthropod appendages (Panganiban et  al. 1997, Williams et  al. 2002). Although it has been difficult to disentangle these broadly overlapping roles of Dll and of other limb-​patterning genes in crustaceans [discussed by Williams (2013)], this section focuses on the role of these genes in the epidermis of crustacean trunk appendages during specification, outgrowth, and segmentation of their main PD axis. Early Dll expression in the round thoracic limb primordia of Drosophila (each comprised of 20–​ 30 cells) is centered on the boundaries of the AP subdivisions (parasegments) of the early embryo (Cohen 1990). Each AP boundary is delineated by adjacent stripes of expression of the segment polarity genes engrailed (en) posteriorly and wingless (wg) anteriorly. The activity of wg, encoding a secreted protein ligand of the Wnt family, plays a key role in Dll activation at the parasegment boundaries and, thus, the induction and correct positioning of the limb primordia along the AP  axis  (Fig.  2.4A) (Cohen et  al. 1993). Furthermore, the lineage-​restricted expression of the homeodomain transcription factor En in the posterior cells provides, from this early stage onward, a widely conserved, lineage-​based mechanism in all arthropods for the subdivision of the developing limbs into anterior and posterior territories (called compartments) (Patel et al. 1989). The correct dorsoventral (DV) positioning of the limb primordia at the ventrolateral sides of the Drosophila embryo is mediated by two signaling pathways: Dll is repressed dorsally by Decapentaplegic (Dpp), a BMP2/​4 ligand of the Transforming growth factor-​β family, and ventrally by Epidermal growth

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Developmental Biology and Larval Ecology factor receptor (Egfr) (Fig. 2.4A) (Goto and Hayashi 1997, Kubota et al. 2000). Although the Dll304 cis-​regulatory element driving early Dll expression in the Drosophila thorax was identified long ago (Vachon et al. 1992), it is still not clear whether the positive and negative inputs from the Wg, Dpp, and Egfr pathways on Dll transcription are direct or not. Fate mapping analysis has shown that each nascent cluster of cells marked by Dll304 activation contributes both to the protopod and telopod of the adult leg, as well as to the adult wing and the rudimentary larval limb of Drosophila known as Keilin’s organ (Fig. 2.4A) (McKay et al. 2009). This is governed by the differential regulation of Dll by other cis-​regulatory elements in subdomains of the nascent cluster when the activity of the early Dll304 decays after a few hours. Dll expression persists in the cells that will give rise to the telopod and the Keilin’s organ, but ceases in the cells that will give rise to the protopod and the dorsal appendage that start expressing other marker genes (Fig. 2.4B) (McKay et al. 2009, Galindo et al. 2011). The cis-​regulatory element that drives Dll expression in about 15 progenitor cells of the telopod receives direct positive inputs from the Wg, Dpp, and Egfr pathways, as well as positive feedback from Dll itself (Estella et al. 2008). The evolutionary conserved zinc-​finger factors of the Sp family, which are expressed early on in the limb primordia across the Pancrustacea, are also required for the maintenance of Dll expression, specification of the ventral appendage fate, and limb growth (Estella and Mann 2010, Schaeper et al. 2010). Overall, the basic subdivision between proximal (Dll-​negative) and distal (Dll-​positive) fates, corresponding to a rudimentary PD axis, is established during Drosophila embryogenesis (Fig.  2.4B). During the early larval stages, the leg disk is divided into two nonoverlapping and antagonistic domains: a distal domain defined by Dll expression and a proximal domain defined by the co-​expression and nuclear localization of the Homothorax (Hth) and Extradenticle (Exd) homeodomain proteins (Fig. 2.4E) (Rieckhof et al. 1997). In the case of Drosophila, these domains of expression coincide with the telopod and protopod regions of the adult leg (Gonzalez-​Crespo and Morata 1996, McKay et al. 2009). Dll Regulation in Direct Developing Crustacean Limbs Expression studies of Dll have been carried out in diverse crustaceans with phyllopodous, biramous, and uniramous appendages (Panganiban et al. 1995, Williams 2013). Early clusters of Dll-​expressing cells appear in the nascent appendage primordia in an anterior-​to-​posterior progression—​first in the anterior head, and later in the gnathal and trunk region. In the most detailed analyses, Dll expression initiates in one or few cells and expands in adjacent cells laterally, anteriorly, and posteriorly (Williams et al. 2002, Hejnol and Scholtz 2004, Prpic 2008). The field of Dll-​expressing cells increases through cell divisions of these Dll-​positive cells and becomes refined to the center of the limb primordium through loss of Dll expression in some peripheral daughter cells. These results suggest that the localized pattern of Dll expression is likely regulated by positional cues (presumably secreted factors) similar to Drosophila. However, unlike in Drosophila, expression becomes confined early to the cells that contribute to the distal region of the developing appendages. Thus, concurrent with their initial specification, crustacean limbs appear to become genetically subdivided into distinct domains of proximal or distal identity, which is also corroborated by more recent lineaging studies (Wolff et al. 2018). Another difference from Drosophila is that these exclusive early domains of Hth/​Exd and Dll expression are not co-​extensive with the protopod and telopod regions in the mature crustacean limbs (Williams et al. 2002, Prpic and Telford 2008). Crustacean limbs are also endowed from inception with the AP patterning information. Using a cross-​reactive antibody, the posterior selector En was found to be expressed in segmentally iterated stripes in crustacean embryos, like in Drosophila and the rest arthropods (Fig. 2.4C) (Patel et al. 1989, Scholtz et al. 1994, Abzhanov and Kaufman 2000c). Dll expression initiates in anterior cells



Crustacean Limb Morphogenesis

abutting the En stripes, but soon afterward expands posteriorly (Hejnol and Scholtz 2004, Browne et al. 2005, Prpic 2008). As a result, appendage primordia straddle the AP compartment boundary and encompass posterior En-​positive and anterior En-​negative cells (Panganiban et al. 1995, Wolff et  al. 2018). Currently, it is not clear how Dll expression initiates in crustaceans and arthropods other than Drosophila; which of the Wg, Dpp, or other pathways are required for limb specification and early PD subdivision; and which Dll cis-​regulatory elements integrate these inputs. The timing and segmental expression of wg are suggestive of its conserved role in Dll activation and early appendage development across the arthropods (Niwa et al. 2000, Prpic et al. 2003, Jockusch et al. 2004, Prpic 2004). However, this role has been functionally confirmed by RNAi knockdown of components of the Wg pathway in holometabolous insects (Grossmann et al. 2009, Ober and Jockusch 2006), but not in hemimetabolous insects (Angelini and Kaufman 2005a). This questions the generality of Wg signaling in appendage specification and early PD patterning in insects, much less pancrustaceans and arthropods in general (Angelini and Kaufman 2005b). In crustaceans, the conserved metameric pattern of wg expression has been demonstrated in two branchiopods (Williams et al. 2002, Prpic 2008, Constantinou et al. 2016), but not in a malacostracan crustacean (Duman-​Scheel et al. 2002). The situation is even less clear with Dpp signaling. In Drosophila, dpp has a dynamic and novel early expression pattern and opposing regulatory functions on Dll during appendage allocation and early patterning (Fig. 2.4A, B) (Cohen et al. 1993, Goto and Hayashi 1997). dpp expression has diverged between Drosophila and other pancrustacean and arthropod embryos, where it is transiently expressed in segmentally transverse stripes and becomes increasingly restricted at the distal tip of the limb bud (Niwa et al. 2000, Prpic et al. 2003, Prpic 2004, Jockusch et al. 2004, Angelini and Kaufman 2005a). Furthermore, dpp knockdown has no effect on Dll expression and early appendage development in a more basal holometabolous insect than Drosophila (Ober and Jockusch 2006). The expression pattern of dpp (but not its function) in crustaceans has been analyzed only in Parhyale (Wolff et al. 2018). Like in other arthropod embryos with direct developing limbs, dpp expression is first detected in segmentally transverse domains corresponding to the dorsal-​fated cells in the limb primordia. High levels of dpp expression persist in a row of anterior dorsal cells abutting the AP boundary in the outgrowing Parhyale limbs, which is reminiscent of dpp expression in the anterior dorsal sector of the Drosophila leg disks. The diverging expression patterns and the paucity of reliable functional data beyond Drosophila, especially for pleiotropic genes such as wg and dpp, make it difficult to reach safe conclusions about the genetic machinery involved in appendage specification and early PD patterning in insects, pancrustaceans, and arthropods. Views vary widely, ranging among (1)  a low conservation in these early mechanisms (Angelini and Kaufman 2005b), (2) the redundancy (or replacement) of the Wg and Dpp signaling activities in early appendage development with (or by) other Wnt and BMP family members in some lineages (Ober and Jockusch 2006), and (3) mechanistic conservation but diverging expression patterns to account for the topological differences between the flat Drosophila leg disc and the three-​dimensional direct limb outgrowths in most other arthropods (Prpic et al. 2003). The limited and fragmentary data also make it challenging to compare stages of appendage development and underlying patterning mechanisms between Drosophila and other pancrustacean lineages. With this caveat in mind, available data suggest remarkable modifications and heterochronic shifts of these mechanisms during the evolutionary transformation of direct into indirect developing limbs. One plausible scenario is that the early embryonic phases of broad Dll activation and later restriction in Drosophila are evolutionary innovations not present in more basal insects and the rest of the pancrustacean lineages. In these lineages, embryos may use patterning mechanisms more comparable to those operating in the early Drosophila leg disk (detailed later) for coupling appendage specification with subdivision into proximal and distal cell fates. Additional evidence

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Developmental Biology and Larval Ecology supporting this scenario comes from the inferred ab initio compartmentalization of the Parhyale limb primordia along the DV axis, in addition to AP, which in Drosophila takes place only during the early second instar larval stage (Wolff et al. 2018). Elaboration of the Limb PD Axis The PD axis of the Drosophila leg matures during larval development as the disk grows to about 10,000 cells. The flat leg disk has a concentric fate map such that cells at the center generate distal structures, whereas cells farther away from the center generate more proximal structures (Fig. 2.4E). Genes specifying the different domains along the PD axis—​referred to as leg gap genes because their mutation results in deletions of the corresponding segments—​are expressed in concentric domains: Dll specifies distal fates in the center, the homeodomain transcription factor dachshund (dac) specifies medial fates in an intermediate ring, the co-​expression of hth/​exd and the zinc-​finger transcription factor teashirt (tsh) specify proximal fates in the disk periphery (Morata 2001, Kojima 2004, Estella et  al. 2012). These expression domains change dynamically during larval development (Fig.  2.4E). Starting with the early juxtaposition of the proximal Hth/​Exd/​Tsh and distal Dll domains, a ring of Dac-​expressing cells arises at an intermediate position between these two domains, followed by a ring of overlapping expression between Dll and Dac, and, finally, an extra proximal ring of cells expressing all leg gap genes. The exclusive and overlapping expression domains of these genes define the mature PD axis that patterns the segmented Drosophila leg (Fig. 2.4F). During the second and early third larval stages, patterning of the leg disk relies on the long-​ range Dpp and Wg morphogens that are activated by the short-​range Hedgehog (Hh) morphogen secreted from the posterior compartment and are expressed close to the AP boundary in anterior dorsal and anterior ventral cells, respectively (Fig. 2.4C) (Basler and Struhl 1994, Diaz-​Benjumea et al. 1994). The discrete domains of leg gap genes are specified by their differential responses to Dpp and Wg signaling, as well as their cross-​regulatory interactions. The central/​distal cells respond to high levels of Dpp and Wg signaling activating Dll (Diaz-​Benjumea et  al. 1994, Lecuit and Cohen 1997). A combination of direct activation by Dll and repression by high Dpp and Wg signaling distally establishes the Dac domain in medial cells (Giorgianni and Mann 2011). Finally, the reciprocal repressive interactions between the hth/​exd/​tsh genes specifying proximal cell fates, and the Wg/​Dpp pathways and their target genes Dll and dac specifying distal and medial cell fates, maintain the subdivision between protopod and telopod (Fig. 2.4C) (Estella et al. 2012). The relative position of the expression domains of the leg gap genes reported in Drosophila are very similar to those observed in direct-​developing stenopodous limbs in crustacean and other arthropod embryos, suggesting that leg gap genes represent a deeply conserved regionalization mechanism also present in nonsegmented onychophoran appendages ( Janssen et al. 2010). Besides Dll expression described earlier, dac, hth, and exd expression has been studied in embryonic limbs of Parhyale and the isopod Porcellio scaber (Abzhanov and Kaufman 2000d, Prpic and Telford 2008, Bruce and Patel 2018). It has been also possible to study the limb phenotypes after CRISPR/​Cas knockout of leg gap genes in Parhyale. The entire telopod is lost after Dll knockout, the proximal telopod after dac knockout, and the protopod and proximal telopod after hth/​exd knockout (Fig. 2.1D) (Kao et al. 2016, Bruce and Patel 2018). Even the phyllopodous limbs of the branchiopod Triops longicaudatus exhibit the three domains of leg gap gene expression along the PD axis (Williams et al. 2002, Sewell et al. 2008). Thus far, the limited and often conflicting functional data outside Drosophila have not been able to resolve the conservation and divergence of the mechanisms setting up the leg gap gene domains in direct-​developing limbs in pancrustacean and other arthropod groups. Unlike the early embryonic lineage-​based compartmentalization of Drosophila limbs along the AP axis, their subdivision along the DV axis takes place later during larval stages and is not based



Crustacean Limb Morphogenesis

on a strict lineage mechanism (Steiner 1976). Although Dpp and Wg synergize in setting up the PD axis (Fig. 2.4C), they act antagonistically in specifying dorsal and ventral cell fates, respectively (Fig. 2.4D) (Svendsen et al. 2015). The “dorsalizing” and “ventralizing” activities of the Dpp and Wg morphogens are mediated by the mutually antagonistic interactions between their downstream T-​box target genes optomotor blind (omb) and Dorsocross (Doc) in the dorsal cells, and the paralogous genes H15/​midline (mid) in the ventral cells (Fig. 2.4D). Cell lineage analyses have recently suggested that the crustacean thoracic limbs of Parhyale are compartmentalized from their inception along both the AP and DV axes (Wolff et al. 2018). This hypothesis is supported by gene expression studies of dorsal and ventral determinants that are detected in the presumptive dorsal and ventral cells from an early stage when Dll starts being expressed in Parhyale limb primordia (Wolff et al. 2018). During the last larval stage of Drosophila, Egfr signaling becomes locally activated in the center of the leg disk and establishes the final pattern in the distal region (Fig. 2.4E) (Galindo et al. 2002, Campbell 2002). A battery of concentrically expressed genes, including the transcription factors aristaless (al), C15, Lim1, Bar (B), apterous (ap), spineless (ss), rotund (rn), and bric-​a-​ brac (bab), specify the pretarsus and the different tarsal subdivisions (Fig. 2.4F) (Kojima 2004, Kojima 2017). The differential activation and repression of these genes by Egfr-​R as signaling, and the extensive positive and negative cross-​regulatory interactions between themselves and with Dll and the more proximally expressed dac, regionalize these most distal parts of the Drosophila leg. Although the roles of Egfr signaling and its downstream targets have not been studied yet in crustaceans, their reported expression in other arthropods and onychophorans suggest they may represent a panarthropod mechanism for distal limb patterning ( Janssen et al. 2010, Grossmann and Prpic 2012). Patterning of Branched Crustacean Appendages Postantennular uniramous limbs evolved independently in several crustacean and arthropod lineages from ancestral branched appendages (Boxshall 2004, Haug et al. 2013, Jockusch 2017). Extant crustaceans uniquely offer the opportunity to compare branched versus unbranched appendage types in a normal developmental context. There are many taxa displaying both uniramous and biramous limbs in their trunk, and taxa in which certain limb branches are present in larval stages but are progressively reduced and eventually lost from adult limbs. We first consider the development of the primary limb axis, i.e. the presence of a single distal element (telopod) in uniramous limbs versus the two distal elements (endopod and exopod) in biramous limbs, and then the development of secondary proximal branches (endites and exites). In Drosophila and other insects normally bearing uniramous legs, duplications of the PD axis can be induced experimentally through surgical grafting experiments or ectopic overlaps of dpp and wg or ectopic Dll expression in the leg disk (Basler and Struhl 1994, Diaz-​Benjumea et al. 1994, Gorfinkiel et al. 1997). Therefore, it was originally proposed that the deployment of the Dpp/​Wg and Dll cassette at different positions in the appendage field may account for the branching pattern of crustacean limbs (Panganiban et al. 1995). The mechanisms behind the bifurcation of the primary limb PD axis have been studied in detail in malacostracan embryos, the amphipod Orchestia, and the isopod Porcellio, which develop uniramous limbs in their pereon and biramous limbs in the pleon (Fig.  2.1C, D) (Hejnol and Scholtz 2004, Wolff and Scholtz 2008). In each species, the early primordia of both limb types are morphologically very similar down to the cellular level. They differentiate during the outgrowth stages when the PD axis remains undivided in uniramous limbs but splits in biramous limbs (Hejnol and Scholtz 2004). Both limb types initiate Dll expression in a contiguous field of cells that is more extended laterally in biramous compared to uniramous limbs. During outgrowth, the

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Developmental Biology and Larval Ecology field of Dll-​expressing cells remains undivided, marking the telopod of uniramous limbs, whereas it splits into two domains marking the endopod and exopod in biramous limbs (Hejnol and Scholtz 2004). This genetic evidence is corroborated by the comparative clonal composition of biramous and uniramous limbs (Wolff and Scholtz 2008) showing that the cells contributing to the endopod and exopod PD axes are contributing to the single PD axis of uniramous limbs (Wolff and Scholtz 2008). Collectively, these data suggest that uniramous limbs in amphipods and isopods are not the product of exopod loss; rather, they are generated by the suppression of the PD subdivision that takes place in biramous limbs. Furthermore, the formation of extra PD axes in these biramous limbs is not the product of extra foci of Wg/​Dpp overlap and Dll activation; rather, it is the product of an expanded contiguous Dll domain and a single limb PD axis splitting into two. It will be interesting to decipher in the future whether a localized downregulation of Dll, cell apoptosis, cell rearrangement, or another morphogenetic mechanism controls limb branching. Studies in Parhyale and Orchestia demonstrate that endites and exites developing on the protopod of amphipod thoracic limbs are secondary branches that develop later in development and from different founder cells than those contributing to the primary limb axis (Wolff et al. 2018, Wolff and Scholtz 2008). Although these proximal outgrowths also express Dll, they are still present after Dll knockout (Kao et al. 2016, Bruce and Patel 2018). Similar results have been obtained for the endites of phyllopodous limbs in branchiopods (Williams et al. 2002, Hiruta et al. 2014) and the endites of pancrustacean gnathal appendages (Browne and Patel 2000, Giorgianni and Patel 2004, Jockusch et al. 2004). Other mechanisms are likely required for crustacean endite and exite formation, with emerging evidence suggesting that these mechanisms resemble the Drosophila wing, rather than leg, genetic hierarchy (Averof and Cohen 1997, Shiga et al. 2017, Clark-​Hachtel and Tomoyasu 2017). Appendage Segmentation Movement of jointed limbs is enabled with flexible intersegmental membranes made of unsclerotized cuticle. Adult Drosophila legs have 10 joints that separate the leg segments (and tarsal subsegments) from each other and from the body wall. Joints are prefigured by patterned gene expression in folds of the leg disk during late larval development, but begin to differentiate a day later during pupal development (Mirth and Akam 2002). These segmental subdivisions occur progressively in the leg disk in a complex order and require the function of the Notch (N) signaling pathway (Fig.  2.4F). Notch signaling establishes the segment boundaries, with the presumptive joints forming at the distal end of each segment, and also promotes leg growth (Rauskolb 2001). The combinatorial action of the leg gap genes and other downstream PD patterning genes localize Notch activation in the joint-​forming concentric rings by controlling the segmental expression of the Notch ligands Delta (Dl) and Serrate (Ser), and the modulator of Notch activity, Fringe (Kojima 2017). In addition to cross-​regulatory interactions among limb-​patterning genes at all levels of the genetic hierarchy, extensive feedback mechanisms with the Notch downstream targets ensure the maintenance and refinement of the joint-​forming regions. A number of genes regulated by Notch function promote the cell shape changes, epithelial invaginations, and cuticle secretion required for proper joint morphogenesis (Suzanne 2016). So far, there are no systematic studies of the role of the Notch pathway in crustacean limb segmentation. Few genes, including the nubbin transcription factor, have been shown to be expressed in iterated rings in segmented stenopodous crustacean limbs (Averof and Cohen 1997, Abzhanov and Kaufman 2000d). Expression and functional studies in representatives of other arthropod subphyla have suggested a conserved role for Notch signaling in limb segment growth and joint formation (Prpic and Damen 2009, Tajiri et al. 2011).



Crustacean Limb Morphogenesis

Limb Diversification along the AP Body Axis Evolution and Function of Hox Genes Crustaceans have attracted the most attention by developmental biologists making the connection between Hox genes and morphological diversity within and across species. Hox genes encode homeodomain transcription factors with a widely conserved role in specifying positional identity along the body axis in bilaterians (Akam 1998, Pearson et al. 2005, Holland 2013). In most arthropods, Hox genes are organized in a single cluster with a relatively loose organization compared to vertebrates (Duboule 2007, Pace et al. 2016). In the best-​reported crustacean assemblies, Hox clusters range from 190 kilobases in the copepod Paracyclopina nana, to 320 to 340 kilobases in daphniids, and more than 2 megabases in Parhyale (Serano et  al. 2016, Kao et  al. 2016, Kim et al. 2018). Comparative studies have suggested that the ancestral arthropod cluster was composed of 10 Hox genes arranged in the following order (Fig.  2.5A): labial (lab), proboscipedia (pb), Hox3, Deformed (Dfd), Sex combs reduced (Scr), fushi tarazu (ftz), Antennapedia (Antp), Ultrabithorax (Ubx), abdominal-​A (abd-​A), and Abdominal-​B (Abd-​B). A few of these Hox genes have been lost in certain arthropod lineages or have acquired new nonhomeotic functions (Hughes and Kaufman 2002). For example, ftz orthologues appear to be absent from malacostracan crustaceans (Serano et al. 2016), while the posteriorly expressed abd-​A orthologues have been specifically degenerated or lost in cirripedes (barnacles) (Deutsch and Mouchel-​Vielh 2003). One of the hallmarks of Hox genes is the correspondence between their physical order in the cluster and their anterior-​to-​posterior progressive expression in nested domains (spatial collinearity), as well as their temporal sequence of activation (temporal collinearity) in some lineages. Temporal collinearity in particular has been proposed as the most likely factor constraining the organization of the Hox genes in an intact and ordered array (Duboule 2007). The most comprehensive expression study in crustaceans has been performed for the nine Hox genes of Parhyale (Serano et al. 2016). Hox genes in Parhyale exhibit both spatial and temporal collinearity; genes closer to one end of the cluster are expressed earlier and in more anterior body segments compared to their neighboring genes located closer to the other end (Fig. 2.5A). Smaller scale expression studies of subsets of these genes (plus ftz, not present in Parhyale) have been carried out in representatives of all major crustacean lineages (Fig. 2.5B and Fig. 2.6) (Shiga et al. 2002, Shiga et al. 2006, Papillon and Telford 2007, Panganiban et al. 1995, Averof and Patel 1997, Abzhanov and Kaufman 1999, Abzhanov and Kaufman 2000b, Abzhanov and Kaufman 2000a, Brena et al. 2005, Vick and Blum 2010, Martin et al. 2016, Deutsch and Mouchel-​Vielh 2003, Fritsch and Richter 2017). Hox genes do not provide the full set of instructions to sculpt morphologically distinct serially homologous appendages. Rather, they modulate the genetic hierarchy that operates in each segment controlling limb development (described earlier) to produce the unique characteristics of each appendage. There is no Hox gene expression in the most anterior head segments developing the first pair of antennae (Hughes and Kaufman 2002). In the absence of Hox input, ventral appendage identity defaults to that of the antenna (Stuart et al. 1991). Hox genes regulate the activity of genes at different levels of the limb-​patterning hierarchy, modifying their regulatory interactions and spatial relationships (Weatherbee et al. 1998, Pavlopoulos and Akam 2011). Furthermore, because Hox genes can exhibit dynamic expression patterns, modulation of genetic hierarchies by Hox genes can take place at different developmental times, producing a whole range of phenotypic changes—​from dramatic to very subtle (Castelli-​Gair Hombria et al. 2016). Although Hox proteins have similar binding preferences, they perform different functions in vivo through the selective activation or repression of downstream target genes. Hox proteins achieve their functional specificity through cooperative protein–​protein interactions with other DNA-​binding partners,

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Developmental Biology and Larval Ecology (A) Abd-B

abd-A

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posterior/later (B) abd-A Amphipoda

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Isopoda

Mysida

Decapoda

Parhyale hawaiensis

Porcellio scaber

Mysidium columbiae* Mysidopsis bahia*

Procambarus clarkii

Fig. 2.5. Hox genes and crustacean appendage diversification (part 1). (A) Schematic representation of the ancestral crustacean Hox cluster. Hox genes have the same transcriptional orientation and exhibit spatial and temporal collinearity. Please refer to the text for gene abbreviations. (B) Expression pattern of abd-​A (gray bars) and Abd-​ B (black bars) in the six biramous pleonic/​abdominal appendages (larger pleopods in light gray and smaller uropods in dark gray) of indicated malacostracan lineages. Abd-​A expression has been analyzed only by the Ubx/​Abd-​A cross-​reactive antibody (Kelsh et al. 1994) in species indicated by asterisks.

including the familiar Exd, Hth, and En proteins, as well as regulatory interactions with other transcription factors, chromatin ,modifiers and effectors of signaling pathways recruited to target the cis-​regulatory elements in a context-​dependent manner (Mann et al. 2009, Rezsohazy et al. 2015). Hox Genes and Crustacean Body Plan and Appendage Diversification Hox gene expression domains coincide well with the subdivisions of the body into morphologically and functionally distinct regions and specialization of the associated appendages. Shifts in the expression boundaries of Hox genes have been associated with macroevolutionary changes in the body organization of pancrustacean lineages. The first evidence came from comparisons between the branchiopod crustacean and insect body plans. The Hox genes Antp, Ubx, and abd-​A are expressed in discrete, nested domains in the stereotyped insect trunk, resulting in the morphological diversification of segments in the thorax and abdomen (Hughes and Kaufman 2002). On the contrary, in situ hybridization studies suggested that all three genes are expressed in largely overlapping domains in the trunk of the branchiopod Artemia franciscana, resulting in uniform segmental and appendage morphologies (Averof and Akam 1995). The basic premise of that study was proved correct. During evolution from ancestors with uniformly segmented bodies, changes in the regulation and expression domains of Hox genes have occurred several times independently in each of the major pancrustacean lineages, giving rise to their unique patterns of segmental specializations (Fig. 2.5B and Fig. 2.6). However, later studies

(A)

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Mystacocarida Derocheilocaris remanei* Mx1

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Anostraca Artemia franciscana Mx1

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Cyclopoida Mesocyclops edax* Mx1

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Cirripedia Sacculina carcini*

Leptostraca Paranebalia belizensis*

* Malacostraca

*

*

*

Isopoda Porcellio scaber

Amphipoda Parhyale hawaiensis

Decapoda Procambarus clarkii

Fig. 2.6. Hox genes and crustacean appendage diversification (part 2). Expression pattern of Scr (light gray), Antp (dark gray), and Ubx (black) in the two maxillary appendages (Mx1 and Mx2) and anterior five trunk appendages (T1–​T5). Strong/​persistent expression is indicated with thick bars, weak/​mosaic expression with thin bars, and temporally increasing/​decreasing expression with ascending/​descending triangles. Protopodal endites and exites are represented with black triangles and ovals, respectively; uniramous limbs with I-​shaped light-​ gray bars; biramous limbs with Y-​shaped light-​gray bars; and phyllopodous limbs with light-​gray bunny shapes. Note there are still some ambiguities about the identity of branches in certain appendages and lineages. In isopods and cirripedes, dark-​gray asterisks denote the expression of Antp messenger RNA that may correspond to bi-​cistronic Ubx/​Antp transcripts that do not produce Antp protein. Ubx expression has been analyzed only by the Ubx/​Abd-​A cross-​reactive antibody (Kelsh et al. 1994) in species indicated by asterisks.

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Developmental Biology and Larval Ecology indicated that only the Ubx protein is actually produced and patterns the uniform branchiopod trunk, raising caution about the limitation of comparative developmental studies that are based solely on in situ hybridization patterns. As revealed in both branchiopod and malacostracan crustaceans, Antp messenger RNA exhibits a much broader distribution than the actual Antp protein as a result of bi-​cistronic Ubx/​Antp read-​through transcripts produced from the Ubx promoter (Shiga et al. 2002, Shiga et al. 2006, Serano et al. 2016). Functional Antp protein is not produced from these bi-​cistronic transcripts, resulting in a more localized Antp activity than that inferred from in situ hybridizations. Similarly, as a result of unresolved silencing mechanisms, the Abd-​A protein is not produced in the trunk epidermis of developing Artemia despite accumulation of the abd-​A transcripts (Hsia et al. 2010). The most solid and striking example has linked the same recurrent change in Hox gene expression to the same appendage modification in the anterior trunk of different crustacean lineages (Fig.  2.6). Maxillipeds (“jaw-​legs”) have a mixed gnathal (feeding) and thoracic (locomotory) identity, and have evolved independently several times. Different crustacean lineages develop any number between zero and three pairs of maxillipeds in their anterior thoracic segments (Averof and Patel 1997). In lineages without maxillipeds, Ubx is expressed in all thoracic segments and it specifies their locomotory identity, whereas Scr is expressed in the more anterior maxillary segments and it specifies gnathal identity. However, in lineages developing one, two, or three maxillipeds, there is a corresponding elimination of Ubx expression in these anterior thoracic segments, accompanied in some cases by the posterior expansion of Scr expression (Fig. 2.6) (Abzhanov and Kaufman 1999, Pavlopoulos et al. 2009, Serano et al. 2016). The suspected causal association between Hox genes and crustacean appendage morphology was demonstrated in gain-​of-​function and loss-​of-​function gene studies for Ubx, Scr, and other Hox genes in Parhyale (Pavlopoulos et al. 2009, Liubicich et al. 2009, Martin et al. 2016). Because of their cross-​regulatory interactions, alterations in Hox gene expression cause spectacular homeotic transformations of one or more appendages into the likeness of a neighboring appendage, thus uncovering their patterning roles. Experimentally induced homeotic transformations in Parhyale revealed that Scr promotes endite development in the protopod and represses telopod development whereas Antp and Ubx repress endite and promote telopod development, specifying the distinct morphologies of clawed gnathopods versus walking legs, respectively (Fig. 2.3A) (Pavlopoulos et al. 2009, Martin et  al. 2016). Considering also the intermediate appendage morphologies obtained in these experimental manipulations, it is reasonable to speculate that the evolutionary homeotic transformations between feeding, hybrid, and thoracic identities have been achieved in various subtle ways through mutations with small effects on the levels and spatiotemporal expression patterns of Scr, Antp, and Ubx (Akam 1998, Averof et al. 2010). Some of these morphological transformations take place during embryonic and postembryonic development of extant lineages, and have, indeed, been associated with the expression dynamics and interactions of these Hox genes (Abzhanov and Kaufman 2000b). Another beautiful illustration of the role of Hox genes in appendage specialization comes from the number of pleopods versus uropods in the abdomen of various malacostracan lineages (Fig. 2.5B). As revealed again by functional studies in Parhyale, the most posteriorly expressed gene Abd-​B promotes the development of biramous limbs in the six abdominal segments, instead of the uniramous limbs present in the thorax (Martin et al. 2016). Its neighboring Hox gene, abd-​A, is co-​ expressed with Abd-​B in the anterior three pairs of pleopods, differentiating them from the posterior three pairs of uropods expressing Abd-​B only (Serano et al. 2016). Unlike amphipods, mysids and isopods and decapods develop five pairs of pleopods and one pair of uropods; this change coincides with a posteriorly extended abd-​A expression domain in these groups (Fig. 2.5B) (Panganiban et al. 1995, Abzhanov and Kaufman 2000a, Abzhanov and Kaufman 2000b, Brena et  al. 2005, Martin et al. 2016). This appendage-​modifying role of abd-​A is also present in the posterior thorax of some



Crustacean Limb Morphogenesis

malacostracans. As demonstrated functionally in Parhyale, the anterior walking limbs (Abd-​A-​ negative) are directed forward whereas the posterior walking limbs (Abd-​A-​positive) are directed backward, giving amphipods their characteristic appearance and name (Martin et al. 2016). A similar function for abd-​A has been proposed in the posterior thorax of the decapod Procambarus, in this case transforming the (Abd-​A-​negative) clawed limbs into (Abd-​A-​positive) walking limbs (Abzhanov and Kaufman 2000b). In an extreme case, the absence of Abd-​A activity from some thecostracan crustaceans, the cirripedes (barnacles), has been associated with their highly reduced abdomen (Deutsch and Mouchel-​Vielh 2003). Changes in the expression domains of Hox genes (through changes in their cis-​regulatory sequences and/​or upstream regulators), changes in the properties of Hox proteins, and changes in the regulation of their downstream target genes have all contributed to the morphological diversity of pancrustacean (and, more generally, animal) body plans and body parts (Hughes and Kaufman 2002, Carroll et al. 2005). In most cases, it remains a big challenge to advance beyond the developmental genes or pathways involved and to identify the actual genetic changes in coding and/​or noncoding sequences responsible for the phenotypic change. One of the landmark exceptions concerns our understanding of the evolution of the limb-​less adult insect abdomen from a crustacean ancestor bearing limbs on all trunk segments (Pavlopoulos and Averof 2002). In Drosophila, the posteriorly expressed Hox genes Ubx, abd-​A, and Abd-​B (but not Antp expressed in the thorax) bind directly and repress the early-​acting Dll304 enhancer, preventing limb specification in the abdomen at a very early stage (Fig. 2.4A) (Vachon et al. 1992, Gebelein et al. 2002). This repression of Dll arose in the insect lineage, because basal hexapods and crustaceans initiate and maintain Dll expression in the presence of posterior Hox proteins in trunk regions with well-​developed appendages (Panganiban et al. 1995, Palopoli and Patel 1998, Lewis et al. 2000, Konopova and Akam 2014). In addition to cis-​ regulatory changes in the Dll locus, rendering it responsive to Hox repression, the evolution of the repressor activity in the insect Ubx protein itself appears to have jointly effected the limb-​less insect abdomen (Galant and Carroll 2002, Ronshaugen et al. 2002). Interestingly, the Artemia Ubx protein (which does not repress Dll) also has a repressor domain, but its repressive activity on Dll is inhibited through intramolecular interactions (Ronshaugen et al. 2002, Taghli-​Lamallem et al. 2008). An outstanding issue concerns the changes in the Abd-​A protein that most likely evolved first (before Ubx) the repressive activity on Dll during insect evolution (Hsia et al. 2010). It should be noted that a similar Dll repressive activity (presumably also modulated by intraprotein interactions) has been mapped in another branchiopod Hox protein, the Daphnia Antp, suppressing the development of comb-​like feeding endites in the first thoracic limb (Fig. 2.3B) (Shiga et al. 2002). In summary, Hox genes represent a widely conserved system that provides axial information to developing cells, and collaborates with the local patterning cues in each segment to control cell fate specification and differentiation. In this role, Hox genes have been a hotspot for evolution to drive the diversification of serially homologous appendages, which is best illustrated by the diversity of pancrustacean appendages within and between species.

CELL AND TISSUE DYNAMICS DRIVING LIMB MORPHOGENESIS Cellular Basis of Crustacean Limb Development: Lessons from Parhyale Limb development has been explained adequately in terms of genes and gene regulatory networks. Much less attention has been given to the cells that are the basic structural and functional units in embryonic tissues producing, integrating, and responding to the physical and chemical mechanisms shaping developing organs. We still need to understand how the information encoded in the genome is translated into the patterned cell activities, such as cell proliferation and death, cell

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Developmental Biology and Larval Ecology rearrangements, and cell shape changes. For example, a major challenge ahead is to understand the cellular basis of serial homology, and how the Hox genes and other limb-​patterning genes modulate these morphogenetic cell behaviors along the AP axis to sculpt appendages with different shapes and sizes. Recent insights into the cellular mechanisms underlying limb bud formation and elongation in direct-​developing arthropod limbs have emerged from integrated studies in Parhyale. Grid-​like Architecture of the Malacostracan Postnaupliar Ectoderm At the time of limb specification, the Parhyale ectoderm is essentially a single-​layered epithelium organized along the AP and DV axes. The ectodermal cells that eventually form the posterior head and entire trunk region (the postnaupliar region) become progressively organized into a series of stereotypic AP rows and mediolateral columns of cells (Dohle et al. 2004, Wolff and Gerberding 2015). Unlike most malacostracan groups that rely on specialized stem-​like cells, called ectoteloblasts, to form this ectodermal grid, amphipods and Parhyale lack ectoteloblasts and rely on the progressive self-​organization of scattered dividing cells into rows in the posterior end of their germ band. The ectodermal grid has the same early metameric organization like the Drosophila and other arthropod embryos, with each new row of cells corresponding to one parasegment (Fig. 2.7A). Initially, each row undergoes two rounds of stereotyped mitotic divisions to generate first a two-​row parasegment and then a four-​row parasegment. The mitotic spindles of these two divisions have a strict longitudinal orientation (parallel to the AP axis), resulting in the arrangement of the daughter cells in highly ordered columns. The different parasegments along the AP axis are indicated by ascending numbers (E1, E2, E3, and so on). In each parasegment, the AP rows of cells are indicated by the letters a, b, c and d, and the mediolateral columns of cells by ascending numbers 0, 1, 2, 3, and so on, with 0 denoting the ventral midline (Fig. 2.7A). The regularity of the ectodermal grid is disrupted by the subsequent more complex but still invariant cell divisions (referred to as differential cleavages) that also contribute to the transition from the parasegmental to the segmental metameric organization of the body. As in the rest arthropods, each segment and the associated appendages are composed of cells from two neighboring parasegments (Fig. 2.7A) (Dohle et al. 2004, Browne et al. 2005, Wolff and Scholtz 2008). For example, the first maxillary segment is formed by parasegments E1 and E2, the second maxillary segment by E2 and E3, the first thoracic segment by E3 and E4, and so on. From anterior to posterior, each segment comprises the posterior descendants of the b row cells and all the descendants of the c and d row cells from one parasegment, as well as all the descendants of the a row cells and the anterior descendants of the b row cells from the following parasegment (Fig. 2.7A). The most medial columns of cells give rise to the nervous system and sternites whereas the more lateral columns of cells give rise to the limbs. For example, the Parhyale second thoracic (T2) limb is composed of the posterior E4b3–​8 cells, all descendants from the E4c3–​9, E4d3–​9, and E5a3–​9 cells, and the anterior E5b3–​9 cells (Fig. 2.7B–​G) (Wolff et al. 2018). A Multilevel Approach to Study Crustacean Limb Development Embryological descriptions, cell labeling experiments, and gene expression studies for Dll have shown that thoracic limbs are specified at the four-​row parasegment stage (Dohle et  al. 2004, Hejnol and Scholtz 2004, Browne et al. 2005). Furthermore, clonal analyses have indicated that the relative AP and DV position of cells in the two-​dimensional grid are also maintained in the mature three-​dimensional amphipod limb (Wolff and Scholtz 2008). These findings have suggested that early patterning mechanisms at the grid stage specify the limb PD axis, and that little, if any, cell rearrangements contribute to limb outgrowth.



Crustacean Limb Morphogenesis (A)

abcd ab cd a b c d

(B)

(G) b E4 c d E5 ab

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0 1 2 34 56 789

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0 123456789

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anterior posterior

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3 4 5 67 89

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Fig. 2.7. Cell lineage restrictions in the Parhyale germ band and developing limbs. (A)  Grid architecture of the germ band with ectodermal cells organized in anterior–​posterior rows and mediolateral columns of cells identified by letters and numbers, respectively. Each parasegmental row of cells undergoes two rounds of longitudinally oriented cell divisions. Cells from two neighboring parasegments contribute to each limb demarcated by the white dotted line. (B–​F) Lateral views of a live-​imaged Parhyale embryo. The highlighted cells from parasegments E4 and E5 give rise to the second thoracic limb. Schematic representations of tracked ectodermal limb cells color-​coded by their (G–​K) anterior–​posterior, (L–​P) dorsal–​ventral, or (Q–​U) proximal–​distal identity. (B–​U) Modified from Wolff et al. (2018).

To address the cellular basis of limb specification and PD outgrowth, a multidisciplinary approach was devised to reconstruct the complete cell lineage of a direct-​developing Parhyale limb and quantify the behaviors of the constituent cells (Wolff et al. 2018). First, transgenic Parhyale embryos with fluorescently labeled nuclei were imaged with multiview light-​sheet fluorescence microscopy over several days of embryogenesis, covering all stages of limb development—​from early specification until late segmentation (Fig. 2.7B–​F). Second, the complete cell lineage of a Parhyale T2 limb was generated based on these multiterabyte and multidimensional image data sets with new open-​source software for cell tracking, called Massive Multi-​view Tracker (or MaMuT) and available as a Fiji/​ImageJ plugin. Third, these accurate and comprehensive reconstructions

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Developmental Biology and Larval Ecology were used to determine the cellular architecture and cellular dynamics operating at the different stages of limb specification, limb bud formation, limb elongation, and segmentation (Fig. 2.7G–​ U). Finally, the deduced cellular models of limb morphogenesis were validated by analyzing the expression patterns of implicated limb-​patterning genes described in the previous section (Wolff et al. 2018). Ab Initio Compartmentalization of the Limb Primordium Lineage tracing experiments have demonstrated the absence of cell mixing between neighboring parasegments (Dohle et al. 2004). The AP compartmentalization of the Parhyale limbs follows the general paradigm of arthropod limbs acquiring the early parasegmental subdivision. In the case of the T2 limb, the anterior compartment consists of En-​negative cells derived from the b, c, and d rows of the E4 parasegment whereas the posterior compartment consists of En-​positive cells derived from the a and b rows of the following E5 parasegment. The AP compartment boundary becomes established at the one-​row stage and is maintained as a straight clonal boundary between these two cell populations throughout limb outgrowth (Fig. 2.7G–​K). Contrary to the AP subdivision, the DV separation in direct-​developing arthropod limbs had remained unexplored until recently. In the case of the Parhyale T2 limbs, the potential lineage restrictions were probed by analyzing the dynamics of digital clones in the live-​imaged and reconstructed limbs. Based on this in silico clonal analysis, a heritable clonal subdivision was inferred between dorsal and ventral compartments from the four-​row parasegment stage onward (Fig. 2.7L–​P). The presumptive DV compartment boundary runs between descendant cells of the E4b and c rows anteriorly, the E5a and b rows posteriorly, and the E4c4–​c5, E4d3–​d4, and E5a4–​a5 cells medially. Some of these predictions were counterintuitive because the neighboring E4b and c rows and the E5a and b rows that will acquire dorsal and ventral fates are first arranged anteroposteriorly in the ectodermal grid. Yet, they were confirmed by analyzing the timing and identity of cells expressing the dorsal and ventral selector genes described in the previous section (Wolff et al. 2018). A single-​cell labeling approach in the early primordia and the study of the end clonal contributions across multiple mature limbs is still needed to provide extra support for this early DV subdivision, and to explore further to what extent it is based solely on a cell lineage mechanism or also on other nonheritable mechanisms. The predicted ab initio AP and DV subdivision of a direct-​developing crustacean limb is very different from the textbook Drosophila paradigm, in which the AP and DV subdivisions take place at different developmental stages during embryonic and larval stages, respectively. In this respect, the Parhyale limb provides much more support to early theoretical models, where the interaction between the different cell populations emerging from the subdivision along the AP and DV axes is setting up the nascent limb PD axis [Boundary Model; (Meinhardt 1983)]. Expression of Dll is first detected at the inferred intersection of the AP and DV boundaries in the four-​row parasegment (Browne et al. 2005, Hejnol and Scholtz 2004, Wolff et al. 2018). During limb outgrowth, the intersection of the AP and DV boundaries marks the tip of the limb bud and persists in this position throughout the later elongation stages (Wolff et al. 2018). Differential Cell Behaviors Contributing to Limb Bud Formation and Elongation The acquisition of spatial coordinates for all constituent epidermal cells and their change over time allowed rigorous quantifications of the cellular mechanisms that remodel the ectodermal epithelium during Parhyale limb outgrowth (Wolff et al. 2018). Furthermore, the identification of the constituent compartments provided a powerful framework to interpret the analyzed cell



Crustacean Limb Morphogenesis

and tissue dynamics. Limb bud formation involves folding of the flat epithelium into a three-​ dimensional bulge (Fig. 2.7). During this transformation, a few cells at the intersection of the compartment boundaries rise above the level of the epithelium, followed by a large-​scale elevation of most dorsal cells and their juxtaposition to the ventral cells that fold underneath, creating the convex surface of the limb bud. As the limb elongates, the original limb bud grows in size and becomes progressively subdivided into an increasing number of partitions (Wolff et al. 2018). What are the cellular mechanisms driving limb outgrowth? So far, two patterned cell behaviors have been identified in Parhyale based on strong correlations that await further experimental testing in the future. First, there is a marked difference in cell proliferation rates between central fast-​dividing cells and peripheral slow-​dividing cells in the limb primordium (Wolff et al. 2018). This finding raises the possibility that spatially controlled cell proliferation may be causally related to limb bud formation. Such a growth-​based morphogenesis model has long been implicated in vertebrate limb outgrowth, and it will be very interesting to explore its generality to invertebrate limb evagination as well. In support of this model, the pharmacological inhibition of cell proliferation in larvae of the branchiopod Artemia results in reduced limb buds (Freeman et al. 1992). During elongation stages, a high concentration of fast-​dividing cells persists at the distal tip of the growing Parhyale limb (at the boundary intersection), which resembles a growth zone that produces much of the new cellular material required for limb outgrowth (Wolff et  al. 2018). Another region of increased proliferation rates is present in the anterior cells abutting the AP boundary. These cells also transduce high levels of Dpp signaling, providing strong correlative evidence for a morphogen-​ dependent control of growth in another arthropod limb besides the Drosophila wing (Restrepo et al. 2014). Second, besides the differential cell proliferation rates, limb elongation is tightly associated and presumably effected by the preferential alignment of mitotic spindles parallel to the PD axis of limb growth (Wolff et al. 2018). Subdivision of the Limb Proximodistal Axis The cellular basis of PD patterning has been addressed indirectly from clonal analyses in Drosophila leg disks (Weigmann and Cohen 1999). In the case of the live-​imaged Parhyale limbs, it has been possible to track individual cells contributing to all the distinct segments backward to the early limb primordium. This analysis shows that the cells that give rise to the proximal, medial, and distal limb segments occupy distinct mediolateral positions in the limb primordium and then distinct PD positions in the early limb bud (Wolff et al. 2018). It will be very interesting in the future to study when and where the proximal, medial, and distal domains of leg gap gene expression are established in the outgrowing Parhyale limbs. This analysis has also revealed a secondary lineage restriction along the PD axis, with no cell mixing observed after the limb bud stage between the cells forming the proximal three limb segments and the distal four segments (Fig. 2.7Q–​U). Unlike the early AP and DV compartmentalizations, these two exclusive PD territories are specified based on their position in the epithelium rather than their lineage. The distal segments originate from a medial disk of cells centered at the intersection of the AP and DV compartment boundaries whereas the proximal segments originate from the peripheral cells (Wolff et al. 2018). Note that the two territories do not correspond to the protopod and telopod subdivisions of the Parhyale limb, but they do overlap with the previously reported co-​ expression of the proximal leg gap genes exd/​hth in the proximal three segments (Prpic and Telford 2008). This correlation, if functionally relevant, contradicts the previously reported Drosophila paradigm in which cells born in the proximal territory readily contribute to more distal leg regions (Weigmann and Cohen 1999).

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Developmental Biology and Larval Ecology Parhyale limb segmentation is a gradual process. Like in Drosophila legs (Rauskolb 2001), segmentation does not occur in a particular order, such as proximal-​to-​distal or distal-​to-​proximal. As the Parhyale limb elongates, it becomes progressively subdivided into an increasing number of units. The first subdivision occurs between the ischium and merus, giving rise to a bipartite limb, with the distal element appearing as a bulge out of the original bud. The ensuing basis/​ischium, propodus/​dactylus, carpus/​propodus, coxa/​basis, and merus/​carpus subdivisions establish the final seven-​segmented pattern (Wolff et al. 2018). It still remains to be investigated whether all segment boundaries represent secondary units of lineage restriction along the limb PD axis, as is the case in the primary ischium/​merus subdivision, and as previously suggested for Drosophila appendage subdivisions (Milan and Cohen 2000). Cellular Basis of Crustacean Appendage Regeneration: Lessons from Parhyale Regeneration is the replacement of damaged or lost tissues or entire body parts in multicellular organisms. The regenerative potential varies widely between different animal groups or even between tissues and developmental stages of the same species (Grillo et al. 2016). Arthropods—​and, in particular, pancrustaceans—​have been long favored as experimental models to study appendage regeneration because they have the capacity to replace missing limbs after predatory attacks, conspecific fights, self-​amputations at predetermined positions after injury (autotomy), or amputations performed by researchers. Malacostracan crustaceans can fully regenerate their limbs throughout their lifetime, and have been used extensively in histological studies of regenerating appendages and in physiological studies of the interplay between regeneration and control of molting by ecdysteroid hormones (Hopkins and Das 2015). However, unlike in insects, much less progress has been made in understanding the molecular genetic mechanisms that drive regenerative growth in crustaceans, and how these mechanisms compare to embryonic appendage development. Another key open issue concerns the identity of progenitor cells and how they are regulated to compensate for the missing limb structures during regeneration in crustaceans. Recent insights into this issue have emerged from studies of limb regeneration in Parhyale described next. Anatomical Aspects of Limb Regeneration The regenerative process after minor injuries, such as damage of the surface cuticle and epidermis, is realized by mitogen-​induced tissue growth (called compensatory regeneration). The repair of the wound involves the replacement of a small amount of tissue during the healing process, without extensive tissue remodeling (Hopkins and Das 2015). More severe injuries, such as the loss of an appendage, require more complex regenerative mechanisms (called epimorphic regeneration), first to seal the gap and then to replace all missing tissues, such as the epidermis, muscles, and nerves (Fig. 2.8A top). Epimorphic regeneration takes place during one or more molting cycles, during which the regenerating appendage grows within the remaining old cuticle. The first step after limb amputation is the sealing of the wound to reestablish a barrier against the external environment. Circulating blood cells (hemocytes) release clotting and melanization factors, and close the wound opening with a plug (a.k.a. scab) (Fig. 2.8A, phase 1). The next step involves the activation of epidermal cells surrounding the wound that change shape and migrate underneath the scab (Fig. 2.8A, phase 2). Epidermal cells form a continuous epithelial layer that further seals the gap, and secrete a thin cuticular layer (known as the cuticle sac, a distinct structure from the new cuticle secreted by the regenerating limb) that protects the growing limb during regeneration (Hopkins and Das 2015). The following period also involves reorganization of existing cells without cell divisions. A mass of cells, called the blastema, forms at the site of the wound and is responsible for replacing



Crustacean Limb Morphogenesis (A) muscle satellite cell

epidermis

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blood cell

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regenerated limb mesodermal muscle lineage satellite cell blood cell

phase 3 Fig. 2.8. Cellular basis of Parhyale limb regeneration. (A) Schematic representation of the cells and tissues in the limb before amputation and at three different phases of regeneration. (B) Contributions of the ectodermal (top) and mesodermal (bottom) lineages to the regenerated limb. Panels reproduced from (Alwes et al. 2016) with permission from the authors.

the missing limb structures through extensive mitotic activity (Hopkins and Das 2015, Alwes et al. 2016). The final step is characterized by extensive cell proliferation, cell death, cell movement, and cell redifferentiation, contributing to the growth and morphogenesis of the new limb segments (Fig. 2.8A, phase 3). Many different cell types, including epithelial cells, muscle precursors, blood cells, and nerve cells, collaborate in rebuilding an intact, fully functional limb that is folded inside the old cuticle, waiting for the next molt to unfold and replace the lost appendage. Cell Lineage Restriction during Parhyale Limb Regeneration Regenerating body parts can be made by pluripotent stem cells, like in planarians, or lineage-​ restricted progenitors from different tissues, like in vertebrates (Tanaka and Reddien 2011). Recent studies in Parhyale have started addressing the identity of cells and their regenerative potential during crustacean limb regeneration (Konstantinides and Averof 2014, Alwes et al. 2016). By taking advantage of the early commitment of cells to distinct germ layers in the highly stereotyped Parhyale embryo, researchers labeled different lineages and followed their contribution to the regenerated limb after amputation. This analysis demonstrated the limited potential of progenitor cells that are restricted by their lineage and proximity to the regenerated appendage (Konstantinides and Averof 2014). The ectodermal lineage contributes exclusively to ectoderm-​derived tissues such as epidermis and neurons (Fig.  2.8B top), whereas the mesodermal lineage contributes exclusively to mesoderm-​derived tissues such as muscles and blood cells (Fig. 2.8B bottom). For a long time, the origin of the new musculature in regenerating pancrustacean limbs has been unclear. The availability of cell and tissue markers in Parhyale enabled the identification of a stem cell-​like population of mesodermal origin physically associated with muscles in the limb (Fig.  2.8B bottom)

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Developmental Biology and Larval Ecology (Konstantinides and Averof 2014). These Pax3/​7-​positive cells serve as progenitors for new muscle fibers during limb regeneration, suggesting they are functionally equivalent or even homologous to vertebrate muscle satellite cells, which are also involved in muscle repair, growth, and regeneration. A major recent breakthrough has been the ability to visualize continuously, with confocal live imaging, regenerating Parhyale appendages with cellular resolution for several days after amputation (Alwes et al. 2016). These recordings have been used to begin reconstructing detailed fate maps of the blastema and track cells in action during regeneration. Cells in the blastema are organized in an outer epithelial layer of ectodermal cells and an inner mass of loosely arranged mesodermal and ectodermal cells. Unlike muscle replacement by satellite cells, there are no specialized stem cells for the epidermis. Most outer ectodermal cells, regardless of their location in the blastema, contribute as progenitors to the epidermis of the new segments by cell proliferation and acquisition of new positional values along the regenerating PD axis (Alwes et al. 2016). Despite the major morphogenetic changes taking place during the transformation of the blastema into a segmented limb, the relative position of epidermal cells along the growing PD axis is largely preserved. The regenerating nerves extend dynamic projections into the blastema and grow into the newly formed segments, restoring the motor input to the new limb (Alwes et al. 2016).

CONCLUSIONS AND FUTURE DIRECTIONS Developmental studies of crustacean appendages are increasingly making research headlines. For instance, very few, if any, developmental biologists could have envisaged just a few years ago that a major discovery, such as invertebrate satellite cells, would be made first in the 15-​year old Parhyale model (Konstantinides and Averof 2014), and only afterward in the Drosophila model that has been scrutinized for more than a century; or that the resolution and completeness of the fate maps of crustacean embryos and appendages could match or even surpass those available for the best classic animal models (Alwes et al. 2011, Alwes et al. 2016, Wolff et al. 2018). All these recent advancements will enable researchers to address fundamental gaps in our knowledge, described throughout this chapter, about crustacean limb specification, patterning, and morphogenesis events; and compare these processes among crustacean, arthropod, and bilaterian lineages with direct-​developing limbs. This inquiry, combined with the crustacean regeneration research, which also has tremendous momentum, is expected to deliver significant contributions to our understanding of how embryonic developmental programs can be redeployed again and again during an animal’s lifetime to replace not only missing limbs, but also adult brain and germ cells (Grillo et al. 2016, Stamataki and Pavlopoulos 2016).

ACKNOWLEDGMENTS We apologize to the authors of numerous research articles who could not be cited because of space limitations. We thank Martin Thiel, Klaus Anger, and Steffen Harzsch for the invitation to contribute to this volume and their extreme patience during the preparation of this chapter. Andy Sombke (Greifswald University) and Stefan Richter (Rostock University) are greatly acknowledged for their kind help with the microcomputed tomographic scans, Pavel Tomancak and Peter Pitrone (MPI-​CBG, Dresden) for their great help with the initial setup of the OpenSPIM microscope, Hajime Watanabe (Osaka University) for providing the transgenic Daphnia magna line, and Bill Lemon for comments on the manuscript. C.W. was supported by the Einstein Foundation Berlin. A.P. was supported by the Howard Hughes Medical Institute.



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Shiga, Y., K. Sagawa, R. Takai, H. Sakaguchi, H. Yamagata, and S. Hayashi. 2006. Transcriptional readthrough of Hox genes Ubx and Antp and their divergent post-​transcriptional control during crustacean evolution. Evolution and Development 8:407–​414. Shiga, Y., R. Yasumoto, H. Yamagata, and S. Hayashi. 2002. Evolving role of Antennapedia protein in arthropod limb patterning. Development 129:3555–​3561. Stamataki, E., and A. Pavlopoulos. 2016. Non-​insect crustacean models in developmental genetics including an encomium to Parhyale hawaiensis. Current Opinion in Genetics and Development 39:149–​156. Steiner, E. 1976. Establishment of compartments in the developing leg imaginal discs of Drosophila melanogaster. Wilhelm Roux Archive of Developmental Biology 180:9–​30. Stuart, J.J., S.J. Brown, R.W. Beeman, and R.E. Denell. 1991. A deficiency of the homeotic complex of the beetle Tribolium. Nature 350:72–​74. Suzanne, M. 2016. Molecular and cellular mechanisms involved in leg joint morphogenesis. Seminars in Cell and Developmental Biology 55:131–​138. Svendsen, P.C., J.R. Ryu, and W.J. Brook. 2015. The expression of the T-​box selector gene midline in the leg imaginal disc is controlled by both transcriptional regulation and cell lineage. Biology Open 4:1707–​1714. Taghli-​Lamallem, O., C. Hsia, M. Ronshaugen, and W. Mcginnis. 2008. Context-​dependent regulation of Hox protein functions by CK2 phosphorylation sites. Development Genes and Evolution 218:321–​332. Tajiri, R., K. Misaki, S. Yonemura, and S. Hayashi. 2011. Joint morphology in the insect leg: evolutionary history inferred from Notch loss-​of-​f unction phenotypes in Drosophila. Development 138:4621–​4626. Tanaka, E.M., and P.W. Reddien. 2011. The cellular basis for animal regeneration. Developmental Cell 21:172–​185. Toyota, K., C. Hiruta, Y. Ogino, S. Miyagawa, T. Okamura, Y. Onishi, N. Tatarazako, and T. Iguchi. 2016. Comparative developmental staging of female and male water fleas Daphnia pulex and Daphnia magna during embryogenesis. Zoological Science 33:31–​37. Ungerer, P., and C. Wolff. 2005. External morphology of limb development in the amphipod Orchestia cavimana (Crustacea, Malacostraca, Peracarida). Zoomorphology 124:89–​99. Vachon, G., B. Cohen, C. Pfeifle, M.E. Mcguffin, J. Botas, and S.M. Cohen. 1992. Homeotic genes of the Bithorax complex repress limb development in the abdomen of the Drosophila embryo through the target gene Distal-​less. Cell 71:437–​450. Vick, P., and M. Blum. 2010. The isopod Asellus aquaticus: a novel arthropod model organism to study evolution of segment identity and patterning. Palaeodiversity 3:89–​97. von Kalm, L., D. Fristrom, and J. Fristrom. 1995. The making of a fly leg: a model for epithelial morphogenesis. Bioessays 17:693–​702. Weatherbee, S.D., G. Halder, J. Kim, A. Hudson, and S. Carroll. 1998. Ultrabithorax regulates genes at several levels of the wing-​patterning hierarchy to shape the development of the Drosophila haltere. Genes and Development 12:1474–​1482. Weigmann, K., and S.M. Cohen. 1999. Lineage-​tracing cells born in different domains along the PD axis of the developing Drosophila leg. Development 126:3823–​3830. Williams, T.A. 2013. Mechanisms of limb patterning in crustaceans. Pages 74–102 in L. Watling and M. Thiel, editors. The Natural History of the Crustacea, Volume 1. Oxford University Press, New York, New York. Williams, T., C. Nulsen, and L.M. Nagy. 2002. A complex role for distal-​less in crustacean appendage development. Developmental Biology 241:302–​312. Wolff, C. 2009. The embryonic development of the malacostracan crustacean Porcellio scaber (Isopoda, Oniscidea). Development Genes and Evolution 219:545–​564. Wolff, C., and M. Gerberding. 2015. “Crustacea”: comparative aspects of early development. Pages 39–​61 in A. Wanninger, editor. Evolutionary Developmental Biology of Invertebrates, Volume 4. Ecdysozoa II: Crustacea. Springer Vienna, Vienna, Austria. Wolff, C., and G. Scholtz. 2002. Cell lineage, axis formation, and the origin of germ layers in the amphipod crustacean Orchestia cavimana. Developmental Biology 250:44–​58.

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3 ORGANOGENESIS

Günther Loose, Günter Vogt, Mireille Charmantier-​Daures, Guy Charmantier, and Steffen Harzsch

Abstract This chapter reviews the development of the major organ systems in crustaceans, including musculature, nervous system, circulatory system, digestive system, osmoregulatory system, excretory system, reproductive system, and sensory organs. It describes the morphological unfolding of these organ systems, which generally follows cleavage, gastrulation, and segmentation in the course of ontogeny. Particular emphasis is given to the organ-​specific temporal dynamics of development, the onset of functionality, and possible correlations with developmental mode, life history, and ecology. The anatomy and cellular characteristics of developing organs are generally better investigated than aspects of physiology, biochemistry, and molecular biology. Investigations in different crustaceans revealed that the speed of development of the various organ systems varies considerably within an individual and between species. As a rule of thumb, anlagen of the nervous tissue, muscular tissue, digestive system, and excretory organs appear first, followed by the circulatory system. Osmoregulatory organs are formed later. The reproductive organs are the last to emerge and to become functional. The mode of development, behavior, and ecology of the postembryonic stages seem to be major determinants that influence the speed differences of organogenesis. This is reflected by timing differences in development of the digestive system between directly and indirectly developing representatives or species with or without lecithotrophic larvae. Other features of the dynamics of organogenesis suggest evolutionary constraints, such as the delayed development of the nervous system in postnaupliar, relative to naupliar, segments in some species. Mechanistic constraints may be involved in heart development and development of nontransitory osmoregulatory organs.

Developmental Biology and Larval Ecology. Edited by Klaus Anger, Steffen Harzsch, and Martin Thiel. © 2020 Oxford University Press. Published 2020 by Oxford University Press.

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INTRODUCTION Organogenesis (the formation and differentiation of the body’s organs and organ systems) embraces a set of developmental processes that, in the crustacean embryo, commonly follow the formation of the germ band. During organogenesis, the cellular material of the previously established germ layers, ectoderm, mesoderm, and endoderm, grows, rearranges, and interacts to form organs by the processes of cell proliferation, tissue folding, cell migration, and condensation. Furthermore, a fundamental part of organogenesis is the differentiation of organ-​specific cell types (e.g., neurons) from previously undifferentiated cells. Organs can be highly complex functional entities, composed of multiple tissue types and often interconnected with different parts of the body; thus, it is more appropriate to speak of organ systems. Spatial delineation of organs is difficult, as the muscular system, for example, contains not only skeletal muscles, but also those that are part of the digestive or reproductive systems. In this chapter, we discuss the development of organ systems selected by their function (e.g., in the case of the muscle system: generating mechanical force). This categorization allows for some overlap between particular organ systems. In our account, we include the nervous system, sensory organs, musculature, digestive system, excretory system, circulatory system, osmoregulatory system, as well as the male and female reproductive systems. Because extensive developmental studies were carried out with only a few crustacean species, knowledge is not available to the same degree for all groups. Crustaceans in which organogenesis is relatively well studied include the branchiopods Artemia salina and Daphnia magna; the peracarideans Parhyale hawaiensis and Porcellio scaber; and the decapods Homarus gammarus, Cherax destructor, and Procambarus virginalis. Daphnia magna and P.  hawaiensis have gained the status of emerging model organisms of development for which some molecular–​genetic techniques have been established. The temporal delineation of organogenesis from the other phases of ontogeny is not straightforward, because some organogenetic processes (e.g., neurogenesis in the brain) are initiated prior to segment formation, while others (e.g., development of reproductive organs) extend far into adulthood. Also, the temporal development of the organs in an individual is not synchronous, which means that some organs are morphologically and functionally complete much earlier than others. The factors on which timing differences depend will be touched on in our discussion.

CENTRAL NERVOUS SYSTEM Emergence and Mitotic Activity of Neuroblasts Neurogenesis in many malacostracan embryos is driven by the mitotic activity of neuronal stem cells, the neuroblasts, which derive from the ectoteloblasts (reviews: Harzsch 2003, Brenneis et al. 2013, Harzsch et al. 2015, and see Chapter 1 in this volume). The emergence and proliferative activity of these stem cells was analyzed in great detail in crayfish such as Cherax destructor. Neuroblasts in these animals originate through a defined, stereotyped lineage by the differential cleavage of precursor cells within the germ band that were previously generated by ectoteloblasts (Scholtz 1992). In C. destructor, thoracic neuroblasts are arranged in a regular bilateral pattern of several files and columns, amounting to a maximum number of 25 to 30 neuroblasts per hemisegment, plus an additional median neuroblast. They undergo repeated unequal divisions to produce ganglion mother cells. The ganglion mother cells later divide again to give birth to ganglion cells (neurons). It was previously believed that ganglion mother cells in decapods divide only once, as they do in insects.

Organogenesis However, there is now evidence from the American lobster that ganglion mother cells in the embryonic crustacean brain undergo not just one but multiple divisions (Benton and Beltz 2002), and this may also be the case in other crustaceans. Sullivan and MacMillan (2001) have examined neurogenesis in late embryonic stages of C. destructor and showed that, in the thoracic and pleonal ganglia, the neurogenic activity in each neuromere ceases during or before the molt to the developmental stage, in which its segmental appendage is first used in coordinated movements. The dynamics of neuroblast proliferation in other malacostracan crustaceans with different life cycles such as crabs and lobsters was also analyzed (Harzsch et al. 1998). As discussed in Harzsch (2003), the temporal patterns of neurogenesis in the ventral ganglia of decapod crustaceans are intimately related to the development of the segmental appendages and maturation of motor behaviors. In the decapod crustaceans studied so far, it appears that an initial short period during early embryogenesis, in which the number of active neuroblasts reaches a peak of up to 30 per hemiganglion, is followed by a long period drawn out over several weeks or even months during which only a subset of the initial neuroblast complement remains mitotically active. Emergence of the Early Brain Scaffold, Maturation of Transmitter Systems, and Neuroendocrine Centers in the Eyestalks Newly generated neurons mature by the extension of cellular processes, axons, and dendrites, collectively referred to as neurites, and bundles of these neurites fasciculate to form a primordial axonal scaffold of the emerging central nervous system (reviews: Whitington 2004, Harzsch et al. 2015). Using fluorescent probes, the primordial brain in crayfish embryos was shown to emerge almost simultaneously in the naupliar neuromeres protocerebrum, deutocerebrum, tritocerebrum, and the mandibular neuromeres (Fig. 3.1B, C, E, F; Vilpoux et al. 2006). Some other malacostracan taxa share a similar principal layout of the primordial brain such as American lobsters (Helluy et al. 1993), the shrimp Palaemonetes argentinus (Harzsch et  al. 1997), and stomatopods (Fischer and Scholtz 2010), whereas amphipods display some minor modifications (Ungerer et al. 2011b). Embryogenesis in crayfish proceeds via a developmental stage called the egg–​nauplius, a term that denotes an early step of germ band formation and comprises head lobes with paired optic anlagen as well as paired anlagen of antennulae and antennae and the mandibles (review: Scholtz 2000, Jirikowski et al. 2013). The cellular material of the naupliar segments and the associated appendage anlagen appear more or less simultaneously, and there is a distinct developmental gap between the differentiation of the naupliar segments and the following segments from the maxilla one onward (Scholtz 2000), which also mirrors the ontogeny of the nervous system in crayfish embryos (Fig. 3.1C). After this “naupliar brain” has formed in crayfish embryos, there is a certain time lag before the maxilla one primordium and its associated neuronal structures develop, and the more caudal segments follow sequentially in the characteristic anterior–​posterior gradient (see Chapter 1 in this volume). Following the egg–​nauplius stage, the brain develops rapidly, as shown by a high rate of mitotic activity of neuroblasts in the brain (Fig. 3.1A, D; Sintoni et al. 2012) and the elaboration of brain structures as labeled with a marker against synaptic proteins (Fig. 3.1E; Vilpoux et al. 2006). After axogenesis, the expression of certain neurotransmitters is the next step in neuronal maturation. In the embryonic brain and developing ventral nerve cord (discussed later), this process was examined by immunohistochemistry in several crustaceans (reviews: Beltz et al. 1992, Harzsch et al. 2015). Studies on the emergence of pigment-​dispersing hormone-​immunoreactive neurons suggest that, at hatching, lobsters in their eyestalk neuropils of the lateral protocerebrum are equipped with a neuronal substrate equivalent to the neuronal networks found in insect brains, constituting the circadian clock (Fig. 3.1.H; Harzsch et  al. 2009). Furthermore, in the eyestalk

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Fig. 3.1. Embryonic development of the brain of crayfish, crabs, and lobsters. (A)  Embryonic brain of the marbled crayfish at around 40% of embryonic development after a 4-​hour pulse of the s-​phase-​specific mitosis marker bromodeoxyuridine and subsequent immunohistochemical detection (Harzsch, Fabritius-​Vilpoux, Sintoni, unpublished). (B) Developing brain in an embryo of the marbled crayfish around 40% of development labeled with a probe for actin. From Vilpoux et al. (2006). (C) Schematic, idealized model of the crayfish egg–​nauplius brain, which 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. The four neuromeres of the naupliar brain are arranged in a mirror symmetry across the horizontal center of the stomodaeum. From Vilpoux et  al. (2006). (D)  Structure and location of proliferative domains (dark gray) in the lobster brain (Homarus americanus) at 25% of embryonic development (E25%). From Harzsch et al. (1999). Proliferation domains 10 and 9/​11 contribute neurons to adult cell clusters 10 (olfactory projection neurons) and 9/​11 (local olfactory interneurons), respectively. (E), (Eʹ) Whole-​mount embryos of lobster embryos Homarus americanus labeled for synaptic proteins (synapsins) to reveal the embryonic nervous system (Harzsch, Benton, Beltz; unpublished). (F), (Fʹ), (Fʺ) Development of the optic anlagen in embryos of the crayfish Cherax destructor labeled with phalloidin, a probe against actin (F:  E35%; Fʹ:  E60%; Fʺ:  E75%). From Vilpoux et  al. (2006). (G) Whole mounts of the optic stalk–​anlagen of the spider crab Hyas araneus at embryonic midstage III after 4 hours of incubation in bromodeoxyuridine (BrdU) and subsequent immunohistochemical detection to show

Organogenesis anlagen of various decapod crustaceans, the ontogenetic expression of several other hormones and neurotransmitters/​modulators has been examined, especially with respect to the maturation of the medulla terminalis X-​organ/​sinus gland complex (Rotllant et al. 1995). Collectively, the embryonic studies indicate that the lobula/​medulla terminalis complex (the “lateral protocerebrum”) and the medial protocerebrum are the areas where the first neurons appear that synthesize a transmitter. The Ventral Nerve Cord: Neurogenesis in the Postnaupliar Segments During growth of the crayfish embryo and also in many other malacostracans, new segments are added caudally to the segments of the egg–​nauplius by the activity of the ectoteloblasts in the caudal growth zone so that new ganglion anlagen develop sequentially in an anterior–​posterior gradient (see Chapter 1 in this volume). Phalloidin labeling of actin was used to identify the emerging connectives and commissures in the developing neuromeres of the ventral nerve cord of crayfish embryos (Alwes and Scholtz 2006, Vilpoux et al. 2006), and studies of several other representatives of the Malacostraca have shown that, in principle, gangliogenesis follows the same general pattern as in the crayfish (Harzsch et al. 1998, Fischer and Scholtz 2010, Ungerer et al. 2011b). With the exception of the midline and one individually identified neuroblast (Ungerer and Scholtz 2008), the vast majority of neuroblasts and the progeny to which they give birth has not yet been identified individually in decapod crustaceans. Therefore, we do not know in most cases which neurons lay down and pioneer the emerging axonal scaffold of commissures and connectives. However, using intracellular tracing techniques, Whitington et al. (1993) identified certain sets of early differentiating neurons (their genealogy has yet to be established), the outgrowing neurites of which participate in pioneering the axonal scaffold within the neuromeres of the ventral nerve cord. Comparing these pioneer neurons between the woodlouse Porcellio scaber and the crayfish C. destructor showed that some of these cells have homologous counterparts in both species whereas others are unique to either one of them (review: Whitington 2004). The presence of segmentally arranged midline neuroblasts in the developing thoracic ganglia has been reported in several malacostracans including crayfish (C. destructor; Scholtz 1992, Sullivan and Macmillan 2001), crabs (Hyas araneus; Harzsch and Dawirs 1994), lobsters (Homarus americanus; Harzsch et al. 1998), as well as peracarid crustaceans (P. scaber and Orchestia cavimana; Gerberding and Scholtz 1999, 2001). The cell division pattern of midline precursor cells was explored in great detail in the amphipod O. cavimana (Gerberding and Scholtz 1999, 2001). The midline neuroblast of O. cavimana is the first crustacean neuroblast of which the lineage and differentiation of its progeny were elucidated, but such data are lacking for other malacostracans. Currently, the water flea Daphnia magna emerges as a new the proliferation zones (PZ1–​3) associated with the developing optic neuropils. From Harzsch et al. (1999). (H) Pigment-​dispersing hormone-​like immunoreactive structures in the developing lobster brain (Homarus americanus) at E45%. Low-​power overview of the developing eyestalks and median brain; black-​and-​white inverted images, projections of z-​stacks of confocal laser scans (C, Cʹ, Cʺ denote clusters of immunoreactive neuronal somata). From Harzsch et al. (2009). (I) Marbled crayfish. Immunolocalization of histamine in an embryonic brain at E60%. From Rieger and Harzsch (2008). A1, 2, antennula and antenna anlagen (in A–​C), antennula and antenna nerves (in D); aMD, anterior portion of the mandibular neuromere; AnN, antenna neuropil; APN, anterior protocerebral neuropil; Ca, Cp, anterior and posterior pre-​esophageal commissures; CON, connective; DC, anlage of deutocerebrum; ES, anlage of esophagus; HE, hemiellipsoid body anlage; ION, inferior esophageal nerve; LG, lamina; LPC, developing lateral protocerebrum; LO, lobula; MD, anlage of mandible (in B, C), mandibular neuromere (in C, D); MDC, mandibular commissure; ME, medulla; MPC, medial protocerebrum; MT, medulla terminalis; MX1, anlage of maxilla one neuromere; MxP1, anlage of maxilliped 1 neuromere; NE, nauplius eye; OA, optic anlagen; OL, olfactory lobe; PEC/​POC, postesophageal commissure; PPN, posterior protocerebral neuropil; PT, protocerebral tract; PZ, proliferation zone; R, developing retina; SON, superior esophageal nerve; ST, stomodaeum; STG, stomatogastric ganglion; TC, tritocerebral neuropil anlage; 10, 9/​11, proliferation domains 10, 9/​11.

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Developmental Biology and Larval Ecology model to analyze molecular aspects of neural stem cell development in crustaceans (Ungerer et al. 2011a, 2012). As in the brain, the subsequent neurogenetic events in the ventral nerve cord are characterized by the ontogenetic acquisition of transmitter phenotypes, a process that was analyzed using immunohistochemical techniques to localize biogenic amines and neuropeptides (review: Harzsch et al. 2015). The functional maturation of the stomatogastric nervous system, with its connections to the ventral nerve cord and also the ontogeny of the pericardial organ, was analyzed in embryonic Homarus gammarus and H. americanus (Casanovas and Meyrand 1995, Pulver and Marder 2002).

SENSORY ORGANS The ommatidia of many adult malacostracan crustaceans contain a group of eight photoreceptors, R1 through R8, that together constitute the rhabdom, where light is absorbed by visual pigments (review: Paulus 2000, Glantz 2014). The photoreceptor axons then project the retinal mosaic topically onto the first optic neuropil, the lamina (Strausfeld and Nässel 1981). Our knowledge on embryonic pattern formation and ommatidial differentiation in crustacean retina (review: Harzsch and Hafner 2006) is derived mostly from studies on crayfish and clawed lobsters (Hafner et al. 1982, Hafner and Tokarski 1998, 2001, Harzsch et al. 1999). The optic primordia in crayfish appear during mid embryogenesis as two fields of tissue located to the left and right of the developing medial brain (Fig. 3.1B, F). The differentiation of the surface epithelium into the retina begins at the lateral sides of the optic primordia and progresses in a medial direction (Hafner and Tokarski 1998). A wave of mitotic activity within a proliferation zone that extends as a band from the anterior to the posterior margins of the optic primordia increases the number of cells in the monolayer of the surface epithelium (Hafner and Tokarski 1998, 2001, Harzsch et al. 1999b). The wave of mitotic activity in the optic primordia is followed by a phase during which the organizational pattern of the cells within the surface epithelium changes in that cells get organized into rows of ommatidial cell clusters, thus forming the patterned portion of the retina. These organizational processes take place in an area called the transition zone and were recognized not only in crayfish, but also in a number of other crustaceans (Fig. 3.1G). New protoommatidia are formed in this region, laterally expanding the patterned region of the retina (Hafner and Tokarski 1998, 2001, Harzsch et al. 1999, Melzer et al. 2000, Wildt and Harzsch 2002). Such comparative studies foster the view that many processes of retinal pattern formation may be evolutionarily conserved across Crustacea into the insects (Melzer et al. 2000, Harzsch and Hafner 2006). In adult decapod crustaceans, the visual input from the compound eyes is mapped onto four columnar optic neuropils—​the lamina, medulla, and the lobula/​lobula plate complex—​which are connected by two successive fiber chiasms (Strausfeld and Nässel 1981). The photoreceptor axons project the retinal mosaic topically onto the first optic neuropil: the lamina. Although the development of the visual neuropils in crustaceans was analyzed in some detail, so far these data are poorly connected to those on retinal formation. However, these studies suggest that, at hatching, malacostracan crustaceans already display a fairly elaborate and functional visual system. It has been demonstrated that the number of facets in the eyes increases massively during larval and adult life—​for example, in rock lobsters (Meyer-​Rochow 1975), anomuran crabs (Eguchi et al. 1989, Meyer-​Rochow et al. 1989), and brachyuran crabs (Harzsch and Dawirs 1996). These findings suggest that, at least in some species of decapod crustaceans, eye formation is a continuous process that persists through embryogenesis well into adult life. Malacostracan Crustacea are equipped with a first (antennulae) and a second (antennae) pair of antennae, associated with the deutocerebrum and tritocerebrum, respectively. The antennulae are equipped with specialized olfactory sensilla (aesthetascs) in addition to bimodal chemo-​and

Organogenesis mechanosensilla, whereas the antennae are equipped only with the latter. The aesthetascs associated with the antennulae house the branched dendrites of olfactory sensory neurons (OSNs), the axons of which target a primary olfactory processing area in the brain (Mellon 2014). The ontogenetic emergence of the aesthetascs and of the associated OSNs has been analyzed in embryos of the Australian crayfish C. destructor (Sandeman and Sandeman 1996, 2003). The primordia of the antennulae emerge very early as part of the appendages of the egg–​nauplius, and their biramous form becomes visible during mid embryogenesis. The first cluster of olfactory sensory neurons appears around 60% of embryonic development in C. destructor and is visible as an aggregation of cells in the distal part of the lateral flagellum, as visualized by Normarsky interference contrast (Sandeman and Sandeman 1996). During subsequent development, more and more clusters of OSNs emerge, sequentially arranged along the annuli of the lateral flagellum, with the most distal clusters being the oldest and the largest. Aesthetasc sensilla lie beneath the surface of the cuticle and are associated with the clusters of OSNs, but the sensilla do not become externalized before postembryonic stage II (POII). At hatching, POI crayfish are virtually bare of any external receptor hairs (Sandeman and Sandeman 1996, Vogt 2008). The anlagen of aesthetasc sensilla are visible beneath the transparent cuticle at the POI stage and become externalized during the molt to the POII stage. During postembryonic life, new annuli and new sensilla are added to the proximal end of the flagella and not at their tips. The newly emerging aesthetasc sensilla become externalized during the molts and crayfish add substantial numbers of segments and aesthetascs to the lateral flagellum during each molt of their postembryonic life (Sandeman and Sandeman 2003).

MUSCULATURE AND NEUROMUSCULAR INNERVATION Cellular Processes of Muscle Formation and Differentiation A considerable body of literature deals with juvenile and adult growth of skeletal muscle fibers, asymmetry in preadult muscle differentiation, transformation of muscle phenotypes, and muscle atrophy in malacostracan crustaceans (see Chapter 5 in Volume 4). The following section focuses on the early development of musculature in Malacostraca, as the majority of publications on crustacean myogenesis are restricted to this group. In Malacostraca mesodermal cell groups underlying the ectodermal appendage buds, the labrum, the head lobes, the telson anlage or that are located around the stomodeum proliferate and give rise to musculature (Manton 1934, Shiino 1942, Weygoldt 1961). It has been shown for the amphipod P. hawaiensis that orthologs of Drosophila premyogenic transcription factors twist and mef2 are expressed in such prospective myogenic cell patches (Price and Patel 2008). The investigations published to date strongly suggest that crustacean muscle development follows the founder cell model, which has been described for insects (review: Schulman et al. 2015) in its major aspects (Harzsch and Kreissl 2010, Jirikowski et al. 2013). Single cell muscle precursors, termed pioneer muscle cells (or founder cells in insects), appear within mesodermal cell clusters. Pioneer muscle cells (Fig. 3.2A, a) show enhanced synthesis of the muscle fiber proteins actin and myosin. Their spindle-​like shape predefines the orientation of the developing muscle. Moreover, these cells can migrate to particular locations within the embryo. The pioneer muscle cell fuses with surrounding mesodermal cells, the fusion-​competent myoblasts, leading to a syncytial muscle precursor (Fig. 3.2B). Myoblast fusion has not yet been directly observed in crustaceans but is likely, considering the absence of cell division in muscle precursors. Within the ­multinucleate precursors, the myofilaments emerge. They are laid down as a nonstriated meshwork of fibrous actin and myosin (Govind 1995, Mykles and Medler 2015), comparable to the fiber

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Fig. 3.2. Crustacean muscle development. (A–​D) Schematic representation of the founder cell model (thick arrows) with variations (thin arrows). (a–​d) Confocal micrographs of crustacean muscle precursors corresponding to the respective stages of the model. (A) Pioneer muscle cell. (B) Fusion of pioneer muscle cell with surrounding mesodermal cells to form a syncytial muscle precursor. (C) The precursor enlarges by further fusion events, forming myofibrils. (D) Maturation of the muscle precursor. (Bʹ) Formation of myofibrils before the initial fusion event. (Cʹ) Myofibril formation after multiple fusion events. (E) Mature myofibrils with the characteristic

Organogenesis arrangement in myocytes of the smooth muscle type in vertebrates. In the next step, the meshwork becomes structured into linear arrays of sarcomeres that give myocytes the characteristic striated appearance (Fig. 3.2C). Furthermore, at this step of myogenesis, the differentiating fast (phasic) and slow (tonic) phenotypes of muscle fibers become observable, which are characterized by short and long sarcomeres, respectively [see references in Mykles and Medler (2015)]. As muscle differentiation proceeds, the myofibrils thicken, condense, and align in parallel fashion (Fig. 3.2D, d). In the mature muscle fiber, the Z-​, A-​, I-​, and H-​bands of the sarcomeres can be detected (Fig. 3.2E). Crustacean muscle development also shows some deviations from the founder cell model. One such deviation lies in the timing of sarcomere formation relative to the time when the muscle precursor becomes multicellular. This aspect of muscle development can vary between individual muscle precursors of the embryo and between species ( Jirikowski 2015). For example, in the freshwater shrimp Neocaridina heteropoda, striation can be observed in single-​cell precursors of extrinsic second-​antenna muscles (Fig. 3.2Bʹ, bʹ). Likewise, cardiomyocyte precursors of H. americanus develop striated filaments without preceding cell fusion (Burrage and Sherman 1979). The mysid Neomysis integer, on the other hand, shows no striation in the multinucleate muscle precursor of the mandible, which at this developmental stage contains no less than 30 nuclei (Fig. 3.2Cʹ, cʹ). Establishment of Neuromuscular Connections Little is known so far about the mechanism guiding formation of neuromuscular connections in crustaceans. In developing trunk musculature of H. americanus, the first synaptic connections are detectable by electron microscopy in multinucleate muscle precursors shortly before the establishment of the sarcomeric banding pattern (Fig. 3.2F) [see references in Mykles and Medler (2015)]. These muscle precursors are innervated by motor neurons located in the corresponding segmental ganglion of the ventral nerve chord. In the insect Drosophila melanogaster, the muscle founder cells provide signals that guide the growth cone of the developing axon to the target site on the muscle surface. Then, innervation, muscle–​tendon attachment, and sarcomerogenesis occur simultaneously (Schulman et al. 2015). The descriptions of lobster neuromuscular development, however, suggest that formation of the synapse precedes sarcomerogenesis (Fig. 3.2G) [see references in Mykles and Medler (2015)]. Electrophysiological studies have shown that the deep abdominal extensor muscles of the lobster embryo have functional innervation already at 33% to 40% of embryonic development (Cole and Lang 1980). Throughout lobster larval life, the innervating nerve fibers proliferate and the area of the synapses increases (Govind et al. 1982, Govind 1995). In the deep pleonal extensor muscle, the innervation field of excitatory and inhibitory neurons is modified from a three-​segmental pattern in early larvae to a segmental pattern in late larvae and juveniles (Stephens and Govind 1981). striated pattern of Z-​, A-​, I-​, and H-​bands. Transmission electron micrograph of a fast muscle fiber of Homarus americanus; after Govind (1995). (a)  Pioneer muscle cell associated with the maxillula in Procambarus virginalis, (b) binucleate extrinsic muscle precursor of antenna, (c) multinucleate extrinsic muscle precursor of antenna, (d) mature muscle in the extensor apparatus of the pleon. (bʹ) Nonfused pioneer muscle cell of extrinsic antenna muscle, Neocaridina heteropoda. (cʹ) Multinucleate mandible muscle precursor, Neomysis integer. (F)  Homarus americanus, basal swimmeret muscle precursors with nerve terminals connecting to the granular sarcoplasm. Scattered myofibrils are present. Modified from Kirk and Govind (1992). (G) Scheme showing muscle precursor innervation sequence. e, excitatory nerve terminal; i, inhibitory nerve terminal; g, granular sarcoplasm; f, myofibrils; F-​Act, stained filamentous actin; M, muscle; MHC, stained myosin (heavy chain); MN, motoneuron; MP, muscle precursor; PMC, pioneer muscle cell. Scale bars: (E) 2 µm, (F) 1 µm, (A–​D, G) 20 µm.

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Developmental Biology and Larval Ecology Topography of Embryonic Musculature In the observed malacostracan species, a muscle precursor pattern is formed that is segmental to a large extent ( Jirikowski et al. 2013) (Fig. 3.3A). As in the insects (Steffens et al. 1995), loss of individual muscle precursors can occur as part of the segment-​specific differentiation process. In every appendage-​bearing hemisegment, transverse muscle precursors are formed laterally and medially to the appendage bud, which give rise to the extrinsic appendage muscles (me and le in Fig. 3.3A). The intrinsic appendage muscles can arise in two different ways: (1) by formation of multinuclear tubular muscle precursors that extend through the appendage rudiment and then subdivide into multiple smaller precursors (Fig. 3.3B), as has been shown for H. americanus (Harzsch and Kreissl 2010); or (2) by formation of pioneer muscle cells at the position of the prospective muscles in the presegmented podomeres (Fig. 3.3C), as in the isopod Porcellio scaber (Kreissl et al. 2008). In the trunk, rudimentary ventral longitudinal muscle precursors give rise to two pairs of continuous muscle strands (Neocaridina heteropoda) (vlm and dlm in Fig. 3.3A). In representatives of Decapoda and Stomatopoda, a longitudinal muscle precursor is also formed posterior to the growth zone (lmp-​post in Fig. 3.3A). This precursor contributes to the continuous ventral longitudinal muscle strands before the embryo is fully segmented. The developmental role of this muscle precursor can be interpreted as providing a mechanism for rapid muscle differentiation in the terminal region of the embryo despite the anteroposterior developmental gradient associated with a short-​germ mode of development. A similar phenomenon is observed in posterior pioneer neurons ( Jirikowski et al. 2015). Muscle precursors other than those of the skeletomusculature also appear at conserved localities, but generally vary more in the way the particular organ musculature is patterned subsequently. A  circular muscle precursor is formed surrounding the developing stomodeum in embryos of N. heteropoda (St in Fig. 3.3A). Precursors of esophageal dilatator muscles are formed from the same muscle precursor and extend in anterolateral and lateral directions. Likewise, circular muscle precursors surrounding the proctodeum are formed (P in Fig. 3.3A, D). In representatives of Decapoda and Stomatopoda, they show a gradual differentiation in an anterior direction along the gut, in opposite direction to the anteroposterior developmental gradient. In P. scaber, muscle precursors associated with the developing midgut caeca form a disk of concentric rings underlying the midgut rudiment (Fig. 3.3E), which are extruded to an array of ring muscles when the midgut rudiments are elongated in the posterior direction. In N. heteropoda (Fig. 3.3F), muscle precursors extend in a bow-​shaped fashion along the sides of the ovoid yolk sac. In Procambarus virginalis, ­star-​shaped muscle precursors spread laterally from the thoracic somites across the surface of the yolk sac in embryonic stage V, forming an irregular meshwork of myocytes (Fig. 3.3G). After stage VIII, irregular contractions of the yolk sac can be observed. The precursors of cardiac muscles in this ­species are arranged in a sheet at embryonic stage VI ( Jirikowski et al. 2010) (Fig. 3.3H). The heart precursor is already formed at stage V and shows contractile activity by then. Sarcomeres are formed in the cardiomyocytes. At stage IX, the myocardium represents a sac, formed from a myocyte ­meshwork. Also, the heart lumen is traversed by muscle fibers (Fig. 3.3I, J). In other representatives, such as amphipods, however, a serial organization is seen in the cardiac muscle precursors (Fig. 3.3K). There, the developing myocytes form a regular helicoid arrangement of muscle fibers around the heart tube as the embryo matures. Cardiomyocytes seem to remain mononucleate (Fig. 3.3L).

DIGESTIVE SYSTEM The digestive system is composed of the food ​collecting and chopping mouthparts and the digestive tract, which is subdivided into the foregut, midgut, and hindgut (Watling 2013). The foregut

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Fig. 3.3. Embryonic muscle precursor patterns. (A) Scheme of Neocaridina heteropoda embryo corresponding to embryonic stage II ( Jirikowski et al 2013), with precursors of intrinsic muscles of antennula and antenna, and precursors of extrinsic muscles in antennula, antenna, and mandible. Longitudinal muscle precursor strands are present posterior to the maxillula segment. The telson anlage shows a posterior longitudinal muscle precursor (lmp-​post), contributing to the ventral longitudinal muscle strand. Additional muscle precursors are located around the stomodeum and the proctodeum. (B) Schematic of intrinsic muscle development in the thoracic endopods of Homarus americanus. Paired tubular muscle precursors elongate within the unsegmented appendage rudiment (left) and enlarge by addition of nuclei in the regions of the prospective podomeres (middle). Then, precursors subdivide into podomere-​specific muscle precursors (right). Modified from

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Developmental Biology and Larval Ecology Fig. 3.3. continued. Harzsch and Kreissl (2010). (C)  Intrinsic appendage muscle development in Porcellio scaber. Formation of paired pioneer muscle cells inside the presegmented podomeres (left). Precursors enlarge (middle) and differentiate at these locations (right). Modified from Kreissl et al. (2008). (D) Stage V embryos of Procambarus virginalis with ring-​like arrangement of pioneer muscle cells around the proctodeum (arrowheads). (E) Porcellio scaber stage 14 embryo with disk of concentric circular pioneer muscle cells (arrowheads) at the tip of the developing left midgut caecum. (F) Late embryos of Neocaridina heteropoda. Muscle precursors form bow-​like extensions across the yolk bulb. (G) Stage V embryos of Procambarus virginalis. Star-​shaped pioneer muscle cells (dotted line) spread across the yolk bulb. (H–​J) Myocardial rudiment of Procambarus virginalis. (H) Stage IV. (I, J) Stage IX. (I) Higher magnification of ( J), showing the mesh-​like fiber organization. (K) Optical section of the myocardial rudiment of Parhyale hawaiensis at 70% developmental time (DT). (L) Projection of the myocardial rudiment at 75% DT. A1, antennula; A2, antenna; dlm, dorsal longitudinal muscle strand; e, eye anlage; F-​Act, labeled filamentous actin; i, intrinsic muscle precursors; le, lateral extrinsic muscle precursors; lmp-​ post, posterior longitudinal muscle precursor; M, myocardium; Md, mandible; me, median extrinsic muscle precursors; MHC, labeled myosin heavy chain; Mx1, maxilla 1; Mx2, maxilla 2; p, proctodeal muscle precursors; st, stomodeal muscle precursors; T, telson anlage; T1–​T3, thoracopods 1 through 3; vlm, ventral longitudinal muscle strand. Arrowheads in (D), (E), (K) and (L) mark positions of nuclei. Scale bars: (D, G, I, L) 20 µm, (E, H) 40 µm, (F) 100 µm.

and hindgut are of ectodermal origin and the midgut and its derivatives are of endodermal origin. The midgut can have voluminous lateral caeca that together form the midgut gland or hepatopancreas. Organogenesis of the digestive system was studied in several crustacean groups, including representatives of the nonmalacostracans and Malacostraca (e.g., Díaz et al. 2008, Štrus et al. 2008, Ponomarenko 2014). In some groups, feeding and digestion starts during the first larval stage already; but, in others, it starts only during the early juvenile phase. The digestive system is relatively simple in nonmalacostracans. Its organogenesis is discussed using the example of the cirriped Austrominius modestus (Ponomarenko 2014). This barnacle develops through nine embryonic, six naupliar, and one cyprid stages. The mouthparts (labrum and mandibles) begin to form in embryonic stage IV. The foregut and hindgut start to develop in stage V from an epidermal invagination on the ventral side of the embryo. In late stage VII, this invagination divides and generates the primordia of foregut and hindgut. The midgut originates from a group of yolky cells, which develop from blastomere 2D. During embryonic stage IX, the yolk is dissolved (Fig. 3.4A) and the former yolk cells divide intensively to form the epithelium of the midgut. This epithelium then fuses with the foregut and hindgut epithelia to form a continuous digestive tract. The mouth is open during this stage but the anus appears only after hatching to the first nauplius stage, which is released into the plankton. Feeding starts in stage II nauplii, which have mandibles with feathered setae suitable for capture of phytoplankton (Foster 1967). The Malacostraca have complex digestive systems with highly diversified mouthparts, a gastric mill for grinding food, and gastric filters for separating the chymus from the solids (Watling 2013). These structures develop and become functional at different times of development, depending on taxon. The white-​striped cleaner shrimp Lysmata amboinensis may serve as an example of the early onset of functionality (Tziouveli et al. 2011). This caridean develops through an embryonic phase, 14 feeding zoeal stages, and a late developmental stage that is morphologically intermediate between a late zoea and an early juvenile. In the literature, this intermediate stage is often referred to as a postlarva (for discussion of terminology, see Chapter 6 in this volume). Well-​developed mouthparts with food sensing and processing structures are already present in the first zoea stage (Fig. 3.4B). Unlike adults, zoeae have no gastric mill, but there are setae in the stomach that help in processing food. The pyloric filters are first visible in zoea IX but are structurally complete only in zoea XIV. These results suggest that the early zoea stages depend on soft food whereas the later zoea stages can handle harder food stuff (Tziouveli et al. 2011). In decapods with a less gradual, more metamorphic, and shorter development, the mouthparts and gastric chewing and filtering structures appear in ­earlier zoeal stages [references in Batel et al. (2014) and Castejón et al. 2015)].

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Fig. 3.4. Organogenesis of the digestive system. (A) Median section of stage IX embryo of the cirriped Austrominius modestus showing primordia of the digestive system. The mouth is already open (arrow). From Ponomarenko (2014), with permission from Humboldt University Berlin. (B) Well-​developed mouthparts of zoea 1 of the shrimp Lysmata amboinensis (the arrow indicates the incisor process of mandible). From Tziouveli et  al. (2011), with permission from Wiley-​Liss. (C–​H) Development of digestive organs in the crayfish Procambarus virginalis. From Vogt (2008), with permission from Wiley-​Liss. (C) Inner margins of mouthparts of a stage III juvenile studded with diverse masticatory and sensory structures. (D) Primordium of the gastric mill in a hatchling. (E) Sclerotized cuticular teeth of the gastric mill in a stage III juvenile (detached from underlying epithelia). (F) Pyloric filter of stage I juvenile showing well-​developed filter tubes (arrow). (G) Cross-​section through the posterior cephalothorax of a hatchling showing the yolk sac, hepatopancreas, and hindgut. (H) Cross-​section of the hepatopancreas tubule of a stage III juvenile displaying structurally differentiated cell types. b, B-​cell; f, F-​cell; fg, foregut; g, ganglion; gc, gill chamber; hg, hindgut; hp, hepatopancreatic tubule system; la, labrum; lt, lateral tooth; lu, lumen; m, mandible; mg, midgut; mt, median tooth; m1, maxilla 1; m2, maxilla 2; pg, paragnath; pp, press plate; r, R-​cell; ys, yolk sac. Scale bars: (A) 50 µm, (B) 10 µm, (C, F, H) 30 µm, (D, E) 40 µm, (G) 200 µm.

The directly developing cambarid freshwater crayfish are good examples of slow development of the digestive organs, and particularly late onset of feeding. After a long lecithotrophic embryonic period, they hatch as juveniles and start feeding during juvenile stage III (Vogt et al. 2004, Seitz et al. 2005). In the marbled crayfish Procambarus virginalis, the primordia of the mouthparts become first visible at about 35% embryonic development (Seitz et al. 2005, Alwes and Scholtz 2006). The first buds to appear are the mandibles, followed successively by the buds of maxillae 1 and 2, and maxillipeds 1 through 3. In hatchlings, the mouthparts are already articulated as in adults, but are devoid of cutting structures and sensory setae. These features appear in stage II juveniles (Vogt 2008), and are well developed and functional only in stage III juveniles, the first feeding stage (Fig. 3.4C).

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Developmental Biology and Larval Ecology The gastric mill is located in the anterior part of the stomach and serves for further grinding of food and mixing of food with digestive enzymes. Its primordium is well recognizable in hatchlings, but the cuticular teeth are not yet developed (Fig. 3.4D). In stage II juveniles, the teeth are more prominent, but only in stage III juveniles are they sufficiently sclerotized and calcified to crush food (Fig. 3.4E). The paired filters in the posterior part of the stomach are already structurally complete in hatchlings. As in the adults, they consist of press plates and several filter tubes covered by setae (Fig. 3.4F). These tubes increase in number with each successive molt (Vogt 2008). The hepatopancreas is the most voluminous organ of the digestive tract of crayfish and it performs intestinal, hepatic, and pancreatic functions (Saborowski 2015). In marbled crayfish, its anlage appears at about 80% embryonic development. In hatchlings, the hepatopancreas consists of a few tubules and collecting ducts (Fig. 3.4G) that terminate at the pyloric filters of the stomach. These tubules increase progressively in number, amounting to several hundred in large adults. The yolk for fueling the embryonic metabolism is located in a separate yolk sac (Fig. 3.4G) and is absorbed and metabolized by the yolk sac epithelium. It is usually exhausted during the second half of juvenile stage III. Spatial separation of the yolk sac and the main digestive tract enables the simultaneous use of yolk and external food, and thus allows gradual active feeding (Vogt 2008). Such a structural separation is probably also the basis of facultative lecithotrophy in many other crustacean larvae and juveniles (Anger and Schuh 1992). The three typical cell types of the hepatopancreas—​the R-​cells (absorption and storage of nutrients), F-​cells (synthesis of digestive enzymes), and B-​cells (presumably synthesis of fat emulsifiers) (Vogt 2002, Saborowski 2015)—​are already discernible in hatchlings, although only the R-​cells seem to be functional at this stage. Although hatchlings do not feed, their R-​cells accumulate increasing amounts of lipid droplets, probably reflecting a transfer of energy carriers from the yolk sac to the hepatopancreas (Vogt 2008). The typical structures of F-​cells and B-​cells are only achieved during stage III juveniles (Fig. 3.4H), indicating full functionality at this life stage. Hammer et al. (2000) measured an increase of activities of the digestive enzymes amylase, trypsin, and nonspecific esterase in early juveniles of red swamp crayfish Procambarus clarkii, confirming that digestive competence is attained at this life stage. In decapods with abbreviated development and feeding larvae, functionality of the hepatopancreas cells is attained earlier—​for instance, in zoea I of the crabs Hyas araneus (Storch and Anger 1983) and Carcinus maenas (Harms et al. 1994).

EXCRETORY ORGANS The excretory organs/​glands of crustaceans have multiple functions, mainly excretory and osmoregulatory through regulation of urine volume and ionic composition. These paired organs derived hypothetically from segmented pairs of excretory organs in ancestral crustaceans. The metameric location of excretory organs and excretory duct openings differs according to the evolutionary position of crustaceans:  (1) maxillary gland in most nonmalacostracans, including branchiopods, ostracods, cirripeds, copepods; (2)  in the lower malacostracans, phyllocarids, hoplocarids, and some peracarids (isopods); and (3)  antennal gland in other peracarids, including amphipods, mysids, and eucarids (euphausids and decapods). In a few species of (1), antennal glands first develop in larvae before their replacement by maxillary glands (discussed later). In adults, excretory glands present a relatively common organization, with three parts:  (1) a coelomic end sac or coelomosac for hemolymph ultrafiltration, (2) an excretory canal or tubule (which may include a labyrinth) for ion reabsorption and other transports, and (3) an exit duct of ectodermal origin sometimes differentiated into a bladder. In most marine and brackish-​water crustaceans, urine is isosmotic to hemolymph. In contrast, some freshwater crustaceans, particularly gammarids and crayfish, produce hypotonic urine,

Organogenesis minimizing ion loss—​a key adaptation to ion-​poor aquatic habitats. The complete ontogeny of excretory glands has been followed in a few species, mainly decapods; in other groups, an excretory gland was observed only in select embryonic or larval stages. In the syncarid Anaspides tasmaniae, Hickman (1936) has pioneered ontogenetic studies of maxillary glands. Mesodermic cells appear around mid development, then the end sac and duct are formed, with a fully developed gland at hatching (Fig. 3.5A–​D). In embryos of the freshwater shrimp Caridina laevis, coelomic sacs develop in the preantennulary, antennulary, and antennal somites (Fig. 3.5E, F). At 30% developmental time (DT), around day 5, two large mesodermic cells occur at the base of the antennae. They start dividing at 70% DT, pointing to a late development of the antennal glands (Nair 1949). In the freshwater Palaemonetes argentinus, antennal glands are present from late embryos at 70% DT as tubules lined by ionocytes containing Na+/​K+-​adenosine triphosphatase (NKA), a structure found in zoeae I through IV. In “decapodids,” the bladder appears, and the distal tubule, proximal tubule, and bladder become distinct in juveniles. All three regions express NKA (Ituarte et al. 2016). The antennal glands of Homarus gammarus develop progressively (Fig. 3.5G–​O). Undifferentiated cells situated near the base of antennae at 23% embryonic DT (egg, meta-​nauplius) form a mesodermal coelomosac at 35% DT and an ectodermic tubular epithelium, with the latter developing into the labyrinth and bladder from 50% embryonic DT to larval stage III. The definite organization of antennal glands of adults is achieved in early juveniles (stages IV and V). NKA, first observed in embryos at 66% DT in the ectodermic sac epithelium, increases quantitatively in the labyrinth (from embryo at 80% DT to larval stage III), then occurs after metamorphosis in the bladder. Excretory organ ontogeny is thus completed after metamorphosis in H. gammarus (Khodabandeh et al. 2006). In contrast, in Astacus leptodactylus, antennal glands develop entirely in the embryo between 43% and 80% DT, when they contain functional ionocytes expressing NKA (Khodabandeh et al. 2005a, b). Hatching juveniles are thus able to produce dilute urine, an essential adaptation to fresh water. Fewer data are available in other groups. Functional excretory antennal glands in larvae, then maxillary glands in adults, are known in cephalocarids, branchiopods, copepods, and amphipods. In cephalocarids, larval antennal glands in the antenna’s proximal segment have the same functional components as adult maxillary glands. They persist in adults, but are small and nonfunctional (Hessler and Elofsson 1992). In branchiopods, functional antennal glands are present in all larval stages. They are present and probably functional in stage II larvae of Artemia (Conte, 1984) or stage IV in Triops longicaudatus (Fryer 1988). They comprise an end sac and an efferent duct with few syncytial cells. In late larvae, the excretory function shifts to well developed maxillary glands (review:  Martin 1992). Amphipods use antennal glands exclusively, which are well described in adult gammarids. The early embryonic development is well studied in Orchestia cavimana (reviews: Scholtz and Wolff 2002, Hunnekuhl and Wolff 2012), but the organogenesis of the antennal glands is poorly documented. Recent embryological studies on Parhyale hawaiensis, if continued, could fill this gap (Browne et al. 2005, Serano et al. 2016).

HEART AND CIRCULATORY SYSTEM Crustaceans have an open circulatory system. In the nonmalacostracans it is relatively simple, but in Malacostraca it is rather complex (Wirkner and Richter 2013). The heart and arteries originate from mesodermal tissue. The heart is among the earliest functional organs in crustaceans. Its appearance is usually closely linked to thoracic segmentation. The brine shrimp Artemia franciscana has one of the simplest circulatory systems in the Crustacea, being composed of the heart, lacunae that channel the hemolymph through the body, and the pericardial cavity that samples the returning blood. The adult heart is a simple tube running

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Organogenesis almost the entire length of the body. It is composed of a single-​layer myocardium lacking endocardium and epicardium, and has segmentally arranged ostial valves and alary ligaments (Økland et al. 1982). Interestingly, only in the posterior section of the heart does the myocardium form a closed tube. In the anterior section, the myocardium forms a trough, the lid of which is provided by the basal lamina of the dorsal epidermis. Heart development begins in the hatching stage from dorsal mesodermal sheets. The heart starts beating during the late nauplius stage (Spicer 1994). With each new segment, a “chamber” with a pair of ostia is added to the heart. When the adult number of segments is reached, the heart grows by elongation of the “chambers.” Ontogeny of the heart in Artemia seems to follow a strict developmental protocol, resulting in a fairly stereotyped pattern of operation not open to the influence of extrinsic factors. Heart rate becomes temperature dependent only after completion of segmentation (Spicer 1994). In the Decapoda, development of the heart starts early as well. In indirect developers, it becomes functional in a late-​embryonic or early-​larval stage, depending on the taxon (McMahon et al. 2002, Huang et al. 2013, Fitzgibbon et al. 2015), whereas in direct developers, which have a relatively long embryonic period but no larval phase, it becomes functional around mid embryogenesis (Alwes and Scholtz 2006). In the shrimp Metapenaeus ensis, which develops through some brooded embryonic, six free naupliar, three protozoal, and three mysis stages, the heart starts beating in the sixth nauplius. Its architecture is still very simple in this stage (Fig. 3.6A, B) (McMahon et al. 2002). Onset of functionality occurs shortly before the shift from lecithotrophy to active feeding. In the following larval stages, the heart differentiates further and several arteries emerge from the anterior and posterior end of the heart, in addition to the initial ophthalmic artery (Fig. 3.6B) (McMahon et al. 2002). In directly developing crayfish, the circulatory system is composed of the heart, the auxiliary heart (cor frontale), numerous arteries and arterioles with distinct walls, hemal sinuses lacking their own epithelia, and the pericard (Vogt 2002). In Procambarus virginalis, these components are well distinguishable in stage IV juveniles—​the first nonlecithotrophic and completely independent life stage (Fig. 3.6C, D). The formation of the myocardium begins in stage V of embryogenesis with the appearance of nonsegmental muscle precursors in the dorsal periphery of the third maxilliped segment ( Jirikowski et al. 2010). In stage VI, which corresponds to roughly 50% embryonic development, the heart primordium is a contractile, trough-​like membranous structure that is dorsally lined by the epidermis, resembling the situation in Artemia. In this stage, irregular contractions can be observed ( Jirikowski et al. 2010). In stage VII, the density of the myofibrillar network has

Fig. 3.5. (A–​D) Ontogeny of excretory organs: Anaspides tasmaniae, ontogeny of the maxillary gland. Mesodermic cells appear around mid development (A, B), then the whole gland is fully developed at hatching (C, D). From Hickman (1936). (E, F) Caridina laevis embryos, ontogeny of the antennal glands. (E) Two mesodermic cells are visible at 30% developmental time (DT). (F) At 70% DT, these two cells divide to form a mass of cells. From Nair (1949). (G–​O) Light micrographs of Homarus gammarus antennal glands, which develop progressively during the embryonic and early postembryonic phases. The coelomosac is formed from mesoderm; the labyrinth and bladder develop from an ectodermic tubular epithelium. The definite organization of the three parts of the antennal gland found in adults is achieved later in early juveniles (stages IV and V), following metamorphosis. (G, H) Embryo at 50% DT. (I) Embryo at 80% DT. ( J) Embryo before hatching. (K) Larval stage I. (L) Larval stage II. (M) Larval stage III. (N, O) Postlarval stage IV. From Khodabandeh et al. (2006), with permission from J. Crust. Biol. ag, antennal gland; ant, antenna; b, bladder; b-​dw, bladder dorsal wall; b-​lu, bladder lumen; b-​vw, bladder ventral wall; cc, coelomosac cavity; cs, coelomosac; csc, coelomosac cells; e, eye; ect, ectoderm; ese, ectodermic sac epithelium; l, labyrinth; lu, lumen; nc, nerve cord; mes, mesoderm; mes-​c, mesoderm cell; mx.2, maxilla 2; mx.g, maxillary gland; mx.g.d, duct of maxillary gland; mx.g.p, external aperture of maxillary gland; mx.g.t, tubule of maxillary gland; mx.t.m, transverse adductor muscle in basal segment of maxilla 2; te, tubular epithelium; up, urinary pore. Scale bars: (G, K–​N) 100 µm, (H) 20 µm, (I) 30 µm, ( J) 40 µm, (O) 80 µm.

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Fig. 3.6. Organogenesis of the circulatory system in Decapoda. (A) Longitudinal section through the posterior thorax of nauplius VI of shrimp Metapenaeus ensis showing location and histology of the heart. From McMahon et al. (2002), with permission from Elsevier. (B) Diagrammatic representation of changes in heart and arteries of M.  ensis during development (the arrow indicates the ostium). After McMahon et  al. (2002), with permission from Elsevier. (C) Transversal section of the posterior cephalothorax of a stage IV juvenile of crayfish Procambarus virginalis showing location and histology of the heart and further components of the circulatory system (the arrow indicates the descending artery). From Vogt et al. (2009), with permission from Wiley-​Liss. (D) Oblique transversal section through the anterior margin of the heart of a stage VII juvenile of Procambarus virginalis showing all of the five valves. From Vogt et al. (2009), with permission from Wiley-​Liss. aa, antennary artery; bc, branchial chamber; bv, branchiopericardial vessel; dt, digestive tract; el, epicard layer of heart; ep, epidermis; g, gills; h, hindgut; ha, hepatopancreatic artery; he, heart; hs, hemal sinus; Ju, juvenile; lu, heart lumen; mu, musculature; M2, second mysis stage; nc, nerve cord; oa, ophthalmic artery; pa, posterior artery; pc, pericard; P1, first protozoea stage; sa, subneural artery; ss, sternal sinus; ts, thoracic sinus. Scale bars: (A) 20 µm, (B, C) 200 µm, (D) 100 µm.

increased and the fibers begin to show cross-​striation. In stage VIII, the dorsal extensions of the contractile network have fused underneath the epidermis, forming a contractile tube. Anteriorly and posteriorly, the myocardium is becoming connected to the anterior and posterior aorta. Between stage VIII and hatching, the ostia become visible. During juvenile development, the myocardium of the heart and the epicardium are continuously thickened. The epicardium is a storage site of glycogen, which can be mobilized on demand and rapidly distributed via the circulation. Likewise, the arterial valves become successively more robust. Now, cross-​striated muscle fibers enable effective modulation of the valve apertures in the five anterior arteries and the posterior artery, facilitating effective channeling of hemolymph to the site of demand. The heart of adult Malacostraca is neurogenically excited by the cardiac ganglion located in the heart wall. This is in contrast to most nonmalacostracans, which have a myogenic heart. Interestingly,

Organogenesis the embryos of decapods seem to have a myogenic heart as well (Harper and Reiber 2001). In freshwater crayfish, the heart becomes neurogenic at the time around hatching. Neurogenic cardiac control is associated with improved functionality and increased environmental sensitivity, which becomes relevant when the juveniles leave the maternal pleopods to start their own life. In embryos of red swamp crayfish Procambarus clarkii, the cardiac output is primarily regulated by changes in heart rate, whereas adults mainly use stroke volume to modulate cardiac output (Harper and Reiber 2006).

ION-​T RANSPORTING EPITHELIA AND OSMOREGULATION In adult crustaceans, osmoregulation is achieved by specialized organs and ion-​transporting cells (ionocytes) (Charmantier et al. 2009, Henry et al. 2012, Lignot and Charmantier 2015) often associated with respiratory organs (gills) or other body parts when gills are absent, particularly in small animals. Their development is key for the ontogeny of osmoregulation (review: Charmantier 1998, Charmantier and Charmantier-​Daures 2001, Charmantier et al. 2009). Excretory organs, also involved in osmoregulation, are discussed earlier in this chapter. Ontogeny with Direct Development Different lineages, such as Peracarida or Cladocera, have independently acquired a mode of brood care in which the embryos are carried in a protective brood pouch by the mother animal. Embryos, osmotically protected in the female body, develop osmoregulatory organs there. Juveniles are able to osmoregulate at hatching, but except for dorsal organs (discussed later), very few data are available on the ontogeny of these osmoregulatory organs during embryogenesis (Charmantier et al. 2009). In embryos of the isopod Cyathura polita, pleopods with exopod buds (later respiratory) and endopods (later osmoregulatory) are externally visible in stage III embryos, the last stage before the manca; they grow further in later stages. Osmoregulatory gills are functional at hatching (Mercer et al. 2013). Gills are the main osmoregulatory organs in adult decapods. External and direct development is limited to few freshwater species. In Astacus leptodactylus, the development of gills bearing functional ionocytes occurs during the embryonic phase and is completed just before hatching (Fig. 3.7A). Gills appear in 65% DT embryos as buds containing undifferentiated cells; branchial cells develop progressively into branchial epithelium where NKA appears in late embryos (90% DT) (Lignot et al. 2005) (Fig. 3.7B). At hatching, juveniles equipped with functional gills are able to hyperosmoregulate and survive in fresh water. Ontogeny with Indirect Development Most decapods have an indirect development. In most species, gills—​not complete and functional at hatching—​develop during larval stages and become functional in juveniles at metamorphosis (Charmantier et  al. 2009, Lignot and Charmantier 2015). For example, in the last zoea (IV) of Carcinus maenas, gills are undifferentiated buds; the first ionocytes appear in gill filaments of megalopae, and fully functional gills exist in juveniles (Cieluch et al. 2004). In some species, ionocytes differentiate during larval stages at extra-​branchial locations before the development of gills, which later take over osmoregulation. This is the case in the thalassinid Lepidophthalmus louisianensis, which osmoregulates through branchiostegites in early larvae, then through gills in later stages (Felder et  al. 1986). Ionocytes differentiate sequentially, first on branchiostegites in

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Fig. 3.7. (A, B) Ontogeny of ion-​transporting epithelia. Embryos of Astacus leptodactylus at 90% development time. From Lignot et  al. (2005), with permission from Springer Verlag. (A)  The short gill filaments show a regular branchial epithelium with a single layer of thick cells. (B)  Fluorescent immunostaining of Na+/​ K+-​ adenosine triphosphatase (NKA) is observed in most gill filaments in all developing gills. Arrows indicate NKA immunostaining. (C) Ontogeny of osmoregulatory epithelia (dotted areas) during postembryonic development of Litopenaeus japonicus. Schematic transverse section of the cephalothorax. From Bouaricha et al. (1994), with permission from Biol. Bull. (D–​H) Immunolocalization of NKA (bright areas) in the branchial cavity of Eriocheir sinensis during postembryonic development. (D) Zoea I: Immunostaining only visible along

Organogenesis larvae, then on epipodites and gills in juveniles and adults of penaeid shrimps (Bouaricha et al. 1994, Pham et al. 2016) (Fig. 3.7C), or on epipodites in larvae, then additionally on branchiostegites of juveniles in homarid lobsters (Lignot and Charmantier 2001). In early zoeae of Eriocheir sinensis, gills are undifferentiated buds; NKA immunostaining is observed in ionocytes along the inner epithelium of branchiostegites. After metamorphosis to megalopa and in juvenile crab stage I, NKA-​immunolabeled ionocytes are located in filaments of posterior (osmoregulatory) gills, not in anterior (respiratory) gills (Cieluch et  al. 2007)  (Fig. 3.7D-H). In different caridean shrimp (Cieluch et al. 2005, Boudour-​Boucheker et al. 2013, 2016, Ituarte et al. 2016), ionocytes are first located on branchiostegites (and sometimes pleurae) in zoeae, on branchiostegites and epipodites in “decapodids” (see Chapter 7, this volume) and early juveniles, then additionally on gills in later juveniles. In species fully adapted to fresh water, such as Macrobrachium pantanalense, expression of the NKA α-​subunit gene (nka-​α) and other transporter genes (V-​H+-​adenosine triphosphatase, Na+/​H+ exchanger) is high in larvae (Boudour-​Boucheker et al. 2016). In Palaemonetes argentinus, nka-​α gene expression is low in early embryos and reaches maximum levels before hatching, as in early larvae in fresh water (Ituarte et al. 2016). Therefore, in decapods, the full development of gills around metamorphosis generally marks the establishment of the adult osmoregulatory capacity. However, in species fully adapted to fresh water, osmoregulation is required at hatching via the acquisition of osmoregulatory function during embryogenesis, either through the development of gills (e.g., in crayfish with direct development) or through extra-​branchial organs (e.g., in freshwater caridean shrimp with indirect development). Small Species In some taxa in which species are small and are often lacking gills (they rely on diffusive tegumental respiration), extra-​branchial osmoregulatory organs occur in adults. For example, ionocytes were identified in the ventral surface of the cephalosome, the swimming legs and integumental windows (copepods), pereopodal disks (amphipods), and epipodites (branchiopods) (review:  Johnson et al. 2014, Gerber et al. 2016). But, the organogenesis of these sites is scarcely described, except for epipodites of branchiopods, which appear as buds on thoracic appendages, just before the limbs swing down to their vertical, adult orientation (Boxshall and Jaume 2009). Dorsal Organs Dorsal organs have been extensively studied, particularly in Artemia sp. since the pioneering studies by Conte et al. (1972) (review: Martin 1992). Various dorsal structures and organs occur in larval or adult crustaceans (branchiopods, copepods, malacostracans)—​named salt gland, dorsal, nuchal,

the inner epithelium of the branchiostegite. (E) Zoea II: Immunostaining in the branchiostegite, but not in gill buds. (F) Zoea V: Gill buds with no immunostaining. (G, H) Megalopa: Positive immunostaining in the three posterior (left) gills, negative in the anterior (right) gills. From Cieluch et al. (2007), with permission of Mar. Ecol. Progr. Ser. (I) Dorsal organ (arrow) in the first instar nauplius larva of Artemia salina (scanning electron microscopy). Note the absence of legs and gills. From Conte (1984), with permission of Academic Press. ( J) Dorsal gland of Gammarus duebeni at embryonic developmental stage V. The organ already present in the early embryo starts degenerating around stage V and disappears by hatching. Degeneration occurs as the coxal gills become visible in the developing embryo. From Morritt and Spicer (1995), with permission from Wiley-​ Liss, Inc. ag, anterior gill; BB, branchial bud; bc, branchial cavity; bst /​BR /​brst, branchiostegite; DO, dorsal organ; E, embryonic eye; EP, epipodite; G, gills (developing coxal gills in J); gb, gill bud; gf, gill filament; gl, gill lamellae; gs, gill shaft; HR, nonbeating heart rudiment; Juv-​Ad, juvenile and adult; lm, lamina; M, mysis; pg, posterior gill; PL, postlarva; PR, pleura; PS, periembryonic space; VM, vitellin membrane; Z = zoea. Scale bars: (A) 100 μm, (B, D–​H) 50 μm.

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Developmental Biology and Larval Ecology or neck organ/​gland; these structures are probably not homologous and may be involved in other functions (review: Martin and Laverack 1992). Here, we consider only dorsal organs with a confirmed osmoregulatory function. In Artemia sp., the cyst already presents a fully functional salt gland. After hatching, in nauplii (Fig. 3.7I), cell differentiation in the dorsal gland begins with the presence of NKA (Sun et al. 1991)  in late nauplii; then, when thoracic epipods develop, the gland is resorbed (Conte 1984). Besides physical forces, caspases promoting apoptosis could be involved in its elimination (Hsieh and Nguyen 2005). Cladoceran embryos have a nuchal gland that disappears after the first postembryonic molt, when epipodites develop as osmoregulatory organs of adults (Aladin and Potts 1995). In early embryos of amphipods, the dorsal gland occurs during the development of the germ layers, and the timing of its degeneration before hatching is species specific (Morritt and Spicer 1995, 1998). In Orchestia sp., the dorsal organ appears before the first ectodermal rows are formed. Its volume increases, then decreases during the last two to three days before hatching, while gills start presenting signs of ion transport (Morrit and Spicer 1996). In Gammarus duebeni, the dorsal organ appears with the germ layer and begins to degenerate in stage V, as coxal gills develop in the embryo (Morritt and Spicer 1995) (Fig. 3.7J). In both species, the shift in the main osmoregulatory organ correlates with changes in osmoregulatory pattern. On land, eggs of the isopod Armadillidium vulgare, which develop in a closed maternal marsupium, have a dorsal organ. Elegant experimentation has shown that this embryonic organ serves in ion regulation, calcium provisioning, and acid excretion before its atrophy in late embryos (Wright and O’Donnell 2010). Artemia: Developmental Biology of Osmoregulatory Organs Little is known about molecular processes underlying the formation and differentiation of osmoregulatory cells and organs. Because of its anatomy, physiology, and availability of its genome, the main model is Artemia’s salt gland. A POU gene APH-​1 was detected in two species (Chavez et al. 1999, Conte 2008, Wang et al. 2012). APH-​1 is expressed before hatching and, in naupliar stages, only in the salt gland. Its expression starts declining in the gland and appears in the thoracic appendages of adults, suggesting its role in the development of the salt gland or similar ion-​transporting epithelia. Trachealess (Af-​trh) was also detected in the salt gland of Artemia nauplii and in thoracic epipods of adults. The two genes may interact to control salt gland differentiation, similar to their interaction controlling tracheal formation in Drosophila (Mitchell and Crews 2002). Future work should investigate these or similar genes in other crustaceans, including decapods with sequential development of osmoregulatory organs.

REPRODUCTIVE SYSTEM The reproductive system is the last of all organ systems to be completed. It is composed of the internally located gonads and gonoducts, and the external accessory sexual organs (Lopez Greco 2013). The gonads are composed of cells of different origin: (1) the germline cells that give rise to the gametes and (2) the stroma cells that form the matrix. The germ line is separated early or late in embryonic development, depending on taxon (Extavour 2005), but the reproductive system as a functioning whole is ready only in adults. The origin and differentiation of the germ line has been investigated in several nonmalacostracan and malacostracan groups, including Anostraca, Cladocera, Copepoda, Amphipoda, Euphausiacea, and Decapoda (references in Ponomarenko 2014). For example, in the whiteleg shrimp Litopenaeus vannamei, the primordial germ cell appears about 285 minutes after spawning as a result of the seventh cell division cycle (Hertzler 2005). Later, this cell divides and the descendants become

Organogenesis embedded in the anlagen of the gonads, where they either develop into oogonia or spermatogonia, depending on the sex. The appearance of the primordial germ cells and their further fate is particularly well investigated in the amphipod Parhyale hawaiensis, a crustacean model of development (Fig. 3.8A, B) (Extavour 2005). In the directly developing marbled crayfish Procambarus virginalis, the paired ovarian anlage is first detectable in hatchlings. It is located underneath the heart and usually contains one or two prominent oogonia per half organ. The oogonia are ensheathed by relatively few matrix cells (Fig. 3.8C) and oviducts are still lacking (Vogt 2007, 2016). In the following juvenile stages, the ovarian envelope becomes thicker and the numbers of oogonia and oocytes increase continuously (Fig. 3.8D). The typical subdivision of the ovary into muscular sheath, interstitium, oogenetic pouches, germaria, and lumen is established from the eighth juvenile stage. Many previtellogenic oocytes are now located in oogenetic pouches delimited by follicle cells. The oogonia are confined to several germaria, which are distributed throughout the ovary (Vogt et al. 2004). In the weeks before first spawning, which occurs around the 16th postembryonic life stage, the oocytes increase dramatically in size as a result of yolk accumulation. The female gonopores and the annulus ventralis (sperm receptacle) in marbled crayfish are formed from juvenile stage IV (Fig. 3.8E) (Vogt et al.

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Fig. 3.8. Organogenesis of female (A–​E) and male (F–​G) reproductive systems. (A) Germ band embryo of amphipod Parhyale hawaiensis with primordial germ cells (arrow) clustered between the head and trunk regions. From Extavour (2005), with permission from Elsevier. (B) Dorsal view of a P. hawaiensis embryo at hatching. The primordial germ cells are now arranged in longitudinal rows (arrowheads) on both body sides. From Extavour (2005), with permission from Elsevier. (C)  Ovarian anlage in hatchling of crayfish Procambarus  virginalis consisting of large oogonium with prominent nucleus and enveloping matrix cells (arrow). The ovarian envelope is not yet closed. (D) Ovary of stage V juvenile characterized by proliferating oocytes (the arrow indicates the follicle cell progenitor). (E) Ventral aspect of a stage V juvenile showing primordia of gonopores (arrows) and annulus ventralis. From Vogt et al. (2004), with permission from Wiley-​Liss. (F) Section of testis anlage of shrimp Litopenaeus vannamei showing single spermatogonia (arrows) surrounded by matrix cells (arrowhead). (G) Testis of crayfish Procambarus clarkii showing cross-​sectioned seminiferous tubules with spermatocytes, spermatozoa (arrow), and Sertoli cells (arrowhead). (F, G) From Krol et  al. (1992), with permission from Wiley-​Liss. av, annulus ventralis; h, head region; hp, hepatopancreas tubule; hs, hemal sinus; n, nucleus; o, oogonium; sc, spermatocytes; t, trunk region. Scale bars: (A, B) 100 µm, (C, D) 10 µm, (E) 100 µm, (F) 20 µm, (G) 50 µm.

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Developmental Biology and Larval Ecology 2004). These structures increase in size during the following juvenile stages but attain their final form only in the mature female. Organogenesis of the male reproductive system is less well investigated. In the red swamp crayfish Procambarus clarkii, the male reproductive organs include the Y-​shaped testis, two vasa deferentia, two gonopores at the coxae of the fifth pereopods, and a pair of copulatory organs (Taketomi et al. 1996). In early stage III juveniles, the testis anlage is small and undifferentiated. It contains a few spermatogonia surrounded by matrix cells (Taketomi et  al. 1996)  as also observed in the whiteleg shrimp Litopenaeus vannamei (Fig. 3.8F). In the following juvenile stages of Procambarus clarkii, seminiferous tubules are formed and the enclosed spermatogonia proliferate. Sperm is produced only in the preadults and adults (Fig. 3.8G). The paired copulatory organs (gonopods) also start to differentiate in stage IV juveniles and are fully developed only in adults.

CONCLUSIONS AND FUTURE DIRECTIONS From our survey of crustacean developmental literature, the following general timing pattern of organogenesis can be derived: Primordials of the nervous tissue, muscular tissue, digestive system, and excretory organs appear first, followed by the circulatory system. Osmoregulatory organs are formed after these events. The reproductive organs are the last to emerge and to become functional. However, the speed of development of the various organ systems can vary between species. Suggested determinants for the speed differences are the general mode of development and the behavior and ecology of the postembryonic stages. Figure 3.9 shows a schematic comparison of the organogenic processes in three selected crustacean genera, with the best data coverage across all organ systems: Artemia, Homarus and Procambarus. The nervous system, muscular system, and digestive system are generally the first to develop, although Artemia and Homarus show some exceptions. The nervous and muscular systems show close functional interconnection to other organ systems. Thus, there may be some mechanistic constraint on their early development. With regard to the development of the nervous system of crayfish, a distinct delay can be observed between the formation of neuronal structures in the anterior three (naupliar) segments of the embryo and the postnaupliar segments—​a feature also observed in other crustacean representatives. This timing pattern has been interpreted as a developmental relic inherited from ancestors, which developed via a nauplius larva. Indirectly developing decapods with feeding larvae (e.g., H. americanus) develop their digestive (and sensory) organs earlier in embryogenesis. In directly developing representatives (e.g., Procambarus), their development is delayed. In species with intermediate abbreviated development, the digestive organs can become functional rather early or rather late, depending on whether they have feeding or lecithotrophic larvae. A link between the speed of organogenesis and lifestyle becomes particularly obvious when the development of excretory organs is compared between species that inhabit water bodies with fluctuations in salinity, salt water, and fresh water. Artemia from hypersaline environments shows earlier development of the excretory organs than Homarus and Procambarus. In both the directly developing crayfish and the indirectly developing lobster, branchial osmoregulatory organs are developed during embryogenesis and become functional just before hatching. In Artemia, the function of the transitory dorsal organ is taken over by the branchial and extra-​ branchial osmoregulatory tissues in the larval phase. Thus, the development of branchial and extra-​ branchial osmoregulatory organs seems to be at least generally characteristic for the postnaupliar phase of development, regardless of the environment (fresh water, saltwater) or whether it occurs in the embryo or in a larva.

Nervous system Musculature Digestive system Gills, Osmoregulatory system Transitory osmoregulatory org. Excretory system Heart/circulatory system Reproductive organs Nervous system Musculature Digestive system Gills, Osmoregulatory system Transitory osmoregulatory org. Excretory system Heart/circulatory system Reproductive organs

hatch

Brood care

Developmental mode

—*

Nervous system Musculature Digestive system Gills, osmoregulatory system Transitory osmoregulatory org. Excretory system Heart/circulatory system Reproductive organs

Eggs carried under pleon

Habitat

Onset of embryogenesis Organ system

Eggs carried under pleon

Indirect development (nauplius larva) indirect development (zoea larva) Direct development

Artemia

Brackish water Salt water

Homarus Procambarus

Fresh water

Genus name

Organogenesis

development

Juvenile

Adult

larval phase

function

Fig. 3.9. Comparison of relative timing of organogenesis in representatives of the genera Artemia, Homarus, and Procambarus. Duration of development and suggested onset of function are displayed for selected organ systems. The onset of function is defined here as first observation of the adult histological properties or, in the case of transitory osmoregulatory organs, histological properties of the functional organ. References for particular species: Artemia sp. (Benesch 1969 for resting eggs, Bartholomaeus et al. 2009, Spicer 1994, Conte 1984). Homarus americanus (Bumpus 1891, Helluy and Beltz 1991, Helluy et al. 1993). Procambarus virginalis (Vogt 2007, 2008, 2016, Alwes and Scholtz 2006, Jirikowski et al. 2010, Vilpoux et al. 2006, Jirikowski et al. 2015, Anderson 1966). Crayfish gill development (Lignot et al. 2005).

In Homarus and Procambarus, the heart begins to beat in mid embryogenesis, whereas in Artemia, it is developed and becomes functional during the larval phase. This difference correlates to mode of development and the size of the egg and hatchling. On the other hand, the speed of heart, and also osmoregulatory organ, development could both reflect a developmental constraint (i.e., the initiation of these organogenetic processes after the naupliar phase of development—​namely, the formation of the anterior three appendage pairs). Common to all crustaceans is the late appearance of the reproductive organs and functionality of the reproductive systems. However, the primordial germ cells can be separated from the soma cells already during early cleavage, depending on taxon. Sperm and eggs are only produced in the preadult and adult life stages. Future work should extend to poorly investigated crustacean groups and to species with ­untypical developmental patterns and environmental adaptations. Combining morphology with biochemistry and ecology is another major future challenge. Of particular interest are the specific functions of transient larval organs and functional shifts in a given organ during development, as demonstrated for regulation of the heart in embryonic and adult crayfish. Identifying the genetic machinery underlying organogenesis is another challenging future issue. Genomic resources are

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Developmental Biology and Larval Ecology increasingly available for crustaceans, as full genome sequences have been established in the past few years for the distantly related crustaceans Daphnia pulex (Colbourne et  al. 2011), Parhyale hawaiensis (Kao et al. 2016), and Procambarus virginalis (Gutekunst et al. 2018). Making use of these genetic databases is expected to advance greatly research on organogenesis in crustaceans.

ACKNOWLEDGMENTS The writing of this chapter was supported by a grant from the DFG Research Training Group 2010 “RESPONSE” to S.H.

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Organogenesis Mitchell, B., and S.T. Crews. 2002. Expression of the Artemia trachealess gene in the salt gland and epipod. Evolution and Development 4:344–​353. Morritt, D., and J.I. Spicer. 1995. Changes in the pattern of osmoregulation in the brackish water amphipod Gammarus duebeni Lilljeborg (Crustacea) during embryonic development. Journal of Experimental Zoology 273:271–​281. Morritt, D., and J.I. Spicer. 1996. Developmental ecophysiology of the beachflea Orchestia gammarellus (Pallas) (Crustacea: Amphipoda): I. Female control of the embryonic environment. Journal of Experimental Marine Biology and Ecology 207:191–​203. Morritt, D., and J.I. Spicer. 1998. The physiological ecology of talitrid amphipods: an update. Canadian Journal of Zoology 76:1965–​1982. Mykles, D.L., and S. Medler. 2015. Skeletal muscle differentiation, growth, and plasticity. Pages 103–​133 In E.S. Chang, and M. Thiel, editors. The Natural History of the Crustacea, Volume 4. Oxford University Press, New York, New York. Nair, K.B. 1949. The embryology of Caridinia laevis Heller. Proceedings of the Indian Academy of Sciences, Section B 29:211–​288. Økland, S., A. Tjønneland, L.N. Larsen, and A. Nylund. 1982. Heart ultrastructure in Branchinecta paludosa, Artemia salina, Branchipus schaefferi, and Streptocephalus sp. (Crustacea, Anostraca). Zoomorphology 101:71–​81. Paulus, H.F. 2000. Phylogeny of the Myriapoda—​Crustacea—​Insecta: a new attempt using photoreceptor structure. Journal of Zoological Systematics and Evolutionary Research 38:189–​208. Pham, D., G. Charmantier, V. Boulo, N. Wabete, D. Ansquer, C. Dauga, E. Grousset, Y. Labreuche, and M. Charmantier-​Daures. 2016. Ontogeny of osmoregulation in the Pacific blue shrimp, Litopenaeus stylirostris (Decapoda, Penaeidae): deciphering the role of the Na+/​K+-​ATPase. Comparative Biochemistry and Physiology B 196–​197:27–​37. Ponomarenko, E.A. 2014. The Embryonic Development of Elminius modestus Darwin, 1854 (Thecostraca: Cirripedia). PhD dissertation. Humboldt University, Berlin, Germany. Price, A.L., and N.H. Patel. 2008. Investigating divergent mechanisms of mesoderm development in arthropods: the expression of Ph-​twist and Ph-​mef2 in Parhyale hawaiensis. Journal of Experimental Zoology B: Molecular and Developmental Evolution 310:24–​40. Pulver, S.R., and E. Marder. 2002. Neuromodulatory complement of the pericardial organs in the embryonic lobster, Homarus americanus. Journal of Comparative Neurology 451:79–​90. Rieger, V., and S. Harzsch. 2008. Embryonic development of the histaminergic system in the ventral nerve cord of the marbled crayfish (Marmorkrebs). Tissue and Cell 40:113–​126. Rotllant, G., M. Charmantier-​Daures, D. De Kleijn, G. Charmantier, and F. Van Herp. 1995. Ontogeny of neuroendocrine centers in the eyestalk of Homarus gammarus embryos: an anatomical and hormonal approach. Invertebrate Reproduction and Development 27:233–​245. Saborowski, R. 2015. Nutrition and digestion. Pages 285–​319 In E.S. Chang, and M. Thiel, editors. The Natural History of the Crustacea, Volume 4: Physiology. Oxford University Press, New York, New York. Sandeman, R.E., and D.C. Sandeman. 1996. Pre-​and postembryonic development, growth and turnover of olfactory receptor neurones in crayfish antennules. Journal of Experimental Biology 199:2409–​2418. Sandeman, R.E., and D.C. Sandeman. 2003 Development, growth, and plasticity in the crayfish olfactory system. Microscopy Research and Technique 60:266–​277. 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. Scholtz, G. 2000. Evolution of the nauplius stage in malacostracan crustaceans. Journal of Zoological Systematics and Evolutionary Research 38:175–​187. Scholtz, G., and C. Wolff. 2002. Cleavage, gastrulation, and germ disc formation of the amphipod Orchestia cavimana (Crustacea, Malacostraca, Peracarida). Contributions to Zoology 71:9–​28. Schulman, V. K., K.C. Dobi, and M. K. Baylies. 2015. Morphogenesis of the somatic musculature in Drosophila melanogaster. Wiley Interdisciplinary Reviews: Developmental Biology 4:313–​334. 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 A 303:393–​405.

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4 DUPLICATED, TWISTED, AND IN THE WRONG PLACE: PATTERNS OF MALFORMATION IN CRUSTACEANS

Gerhard Scholtz

Abstract The study of malformations is an important tool to understand mechanisms and causes of development and regeneration. Moreover, malformations indicate the morphological potential of living beings. Hence, a deeper understanding of how, to what degree, and why organismal structures can deviate from their normal expression is interesting in an evolutionary and ecological context. Like other arthropods, and animals in general, crustaceans show a certain variety of naturally occurring malformations of different body parts. This review is restricted to those that affect the axes of appendages and the trunk. Hence, the various patterns of axis distortion are described and classified. At the general level, malformations concerning limbs are discriminated from those that alter other body outgrowths and those that affect the pattern of the trunk. Among malformation of limbs and other body appendages, misplaced structures, fissions, and fusions are classified. Conjoined twins and distorted body segments are the main features of trunk malformations. The putative causes of malformations are discussed with respect to comparative and experimental approaches. Furthermore, gene expression studies, theories, and models, such as Hans Meinhardt’s Boundary Model, are applied to explain malformations at the level of pattern formation. Apparently, many malformations are not genetic mutations and thus not inheritable, but are instead the result of distortions during early development and regeneration artifacts based on injuries, high temperature, and toxic substances. Compared with other arthropod groups, there are very few experimental studies addressing malformations in crustaceans. Hence, the causes for specific patterns of deformities remain largely obscure.

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INTRODUCTION Malformations in humans, animals, and plants have always fascinated people. This is reflected by numerous mentions, descriptions, and illustrations of such phenomena in early encyclopedic, zoological, and anatomical works (e.g., Seba 1735–​64, Rösel von Rosenhof 1755). Furthermore, malformations were frequently exhibited as objects of amusement in curiosity cabinets (Bredekamp et al. 2000, Mauries 2002, Martin 2010) as well as in museums of natural history and—​nowadays, in particular—​on the Internet (e.g., Langley 2013, Holland 2014). However, apart from mere fascination, these unusual morphologies also offer the opportunity to gain new scientific insights (e.g., Lewis 1994, Blumberg 2009, Janssen 2013, Scholtz and Brenneis 2016). A  comparative analytical approach to abnormal development and the artificial creation of malformations allow us to draw conclusions about the principles of processes and mechanisms of an organism’s form. As is the case for arthropods in general, a number of accounts of naturally occurring malformations in crustaceans can be found in the literature. However, compared with hexapods and chelicerates, much fewer cases have been reported in crustaceans. Furthermore, in contrast to hexapods, crustaceans were rarely chosen as subjects for experimental approaches for creating artificially malformed animals. Nevertheless, starting with Berniz (1671) and Rösel von Rosenhof (1755), a certain number of reports have accumulated in which a variety of patterns have been described. Aberrant structures occur in embryos, larvae, and adult crustaceans. Quite frequently, speculations about the causation of abnormalities have been put forward, such as toxic substances, mutations, injuries, regeneration, inbreeding, and parasites. The few experimental studies on crustacean malformations led to some insights. Nevertheless, the understanding of the mechanisms in terms of morphogenetic gradients, gene expression, and physiology is still in its infancy (see Mittenthal 1980, 1981, Shelton et al. 1981, Nakatani et al. 1998, Meinhardt 2009, Liubicich et al. 2009, Pavlopoulos et al. 2009, Martin et al. 2016). Likewise, the knowledge about external factors, such as toxic substances in the environment, is still sparse, and the few experiments in this direction do not contribute much to an explanation of the specificity of the observed kinds of malformation (e.g., Weis et al. 1992, Sundelin and Eriksson 1998). Moreover, morphological malformations like those treated in this review are different from abnormalities such as lesions and shell diseases that often occur in polluted areas, such as the Gulf of Mexico after the dramatic oil spill in 2010 (Felder et al. 2014). Many of the reports on crustacean malformations deal with absent or irregularly formed body parts, such as missing spines, deformed and asymmetric carapaces and pleons, or shortened and crumbled appendages (e.g., Mantellato et al. 2000, Elmoor-​Loureiro 2004, Fernandes et al. 2010, Araújo and Calado 2012, Miličić et al. 2013, Rasheed et al. 2014). Because these deformities are often just individual cases and show no general pattern, the most probable causes for them are wound healing processes after an injury or molting artifacts and perhaps developmental perturbations based on toxic substances. Another class of frequently observed altered morphologies concern intersexes and gynandromorphs, which show a combination of transformed appendages—​in particular, gonopods and segmental structures such as gonopores (e.g., Farmer 1972, Ahyong and Ng 2008, Rudolph and Verdi 2010, Martin and Scholtz 2012). At least in decapods, these changes are the result of dysfunctional androgenic glands (e.g., Sagi et al. 1990). Parasite infestations cause a third group of malformations. For instance, bopyrid isopods induce lateral swelling of the branchial region in decapods (Williams and Boyko 2012), and rhizocephalan barnacles castrate and feminize male crab hosts, which leads to a transformed pleon and other external structures (Kristensen et al. 2012). The following review focuses on an additional class of malformations that is characterized by disturbances of the axes that make up the body and its parts. More specifically, this means the occurrence of multiplications, fusions, and perturbations in the proximodistal axes of limbs and

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Developmental Biology and Larval Ecology other appendages, and in the anteroposterior axis of the body. Furthermore, appendages that are in the wrong place along the body axis are covered. By far, most cases of observed and described aberrant structures of this kind occur in crustacean limbs. In contrast to this, trunk anomalies are comparatively rare. These relate to asymmetries, to missing or partly fused segments and hypertrophied segments, or to the interesting phenomenon of spiral segmentation. The most dramatic malformations concern the duplication of parts of the longitudinal body axis that leads to the occurrence of conjoined twins. The patterns described in this review are not special for crustaceans. On the contrary, corresponding cases appear in the other major arthropod groups—​namely, chelicerates, myriapods, and hexapods (Bateson 1894, Przibram 1910, 1921). Moreover, misplaced appendages, multiplications of limb parts, and conjoined twins have been reported from most major bilaterian taxa, including vertebrates and humans (Bateson 1894, Korschelt 1907, Przibram 1910, 1921, Blumberg 2009, Lobo et al. 2014, Scholtz et al. 2014). Hence, one can assume some general principles of pattern formation that were already present in the bilaterian stem species. There is a certain taxonomic bias in the reports about crustacean malformation. By far, most data concern decapod species—​in particular, lobsters, crayfishes, and crabs. Some of the documented cases relate to other malacostracan groups and even less to the non-malacostracan crustaceans. This apparently does not mean that decapods are more sensitive to malformations than other crustaceans. Rather, this bias is a result of the size, accessibility, and economic importance of many species of this group. In addition to the insufficient state of knowledge, it has to be stressed that the questions of causes and explanations of malformations are not trivial. First, one has to discriminate between proximate and ultimate causes (see Mayr 1987, Scholtz 2008). The proximate cause is what actually happened to the animal and what triggered the reaction that led to the malformed structure. The ultimate cause is the set of mechanisms that developed in the course of evolution and that shape the reaction based on their abilities and constraints. Accordingly, explanations for an observed malformation are possible at various levels. For instance, a malformation like a bifurcated moveable finger of a crab claw is explainable as the result of an injury during combat with a conspecific. At the same time, it is the product of the capability of the species for wound healing and tissue regeneration that leads to the observed pattern. At the molecular level, the healing process involves interaction and regulation of genes that play a role in morphogenesis. Hence, explanations just stressing pollutants or Hox genes are insufficient. Nevertheless, what can be safely concluded is that most malformations described in the following are caused by chemical and/​or mechanical perturbations during development or regeneration. Hence, they are not mutants at the genetic level.

MALFORMED LIMBS Heteromorphoses or Homoeotic Changes The phenomenon of misplaced body parts has been called heteromorphosis (Loeb 1891)  (Figs. 4.1 and 4.2). A special case of heteromorphosis is the replacement of a serial structure by a corresponding but different structure from another body region—​a phenomenon named homoeosis by Bateson (1894). There are some cases of heteromorphosis and homoeosis reported for crustacean limbs. The documented cases can be classified into three major groups. The first group contains the replacement of serially homologous limbs or limb parts—​in other words, clear cases of homoeosis (Figs. 4.1 and 4.2). On the one hand, this replacement concerns limbs from adjacent segments (Figs. 4.1G and 4.2B). For instance, specimens of the crayfish Astacus leptodactylus show chelate claws in the fourth pereopods resembling those of the third pereopods. In



Patterns of Malformation in Crustaceans

turn, in some cases, the third pereopods are achelate (Zalpeter 1927). A third maxilliped equipped with a reduced version of the claw bearing first pereopods instead of the normal palp was found in four crab species: Cancer pagurus, Chionoecetes opilio, Dromia personata, and Uca rapax (Bateson 1894, Przibram 1910, Motoh and Toyota 2003, Lira et al. 2013) (Fig. 4.1G). Similarly, Needham (1941) described a male isopod Proasellus meridianus in which, additional to the normal equipment, the protopodite of the first pleopod bears a biramous appendage resembling the second pleopod and, to some extent, the uropods. On the other hand, limb structures of distant segments and tagmata have been misplaced (Figs. 4.1E, F and 4.2A). In a specimen of the spiny lobster Panulirus argus, the endopod of a pleopod forms a second antenna flagellum (Fig. 4.1F); in another individual a fourth, pleopod was found, in which a somewhat reduced pereopod replaces the endopod (Fausto-​Filho and Da Costa 1977) (Fig. 4.1E). The second group consists of a limb in a normally limbless segment, but in a position where a limb can be expected. This is a special case of homoeosis. Gordon (1963) reported a male spider crab with pleopods in the third to fifth pleon segments, whereas pleopods do not normally occur in eubrachyurans. All these extra limbs resemble the second pair of pleopods, which are involved in

(A)

(B)

(E)

(D)

(F)

(C)

(G)

(G’)

Fig. 4.1. Heteromorphoses. (A) Antenna-​like structure replaces the left eye of the rock lobster Palinurus elephas. The right eye is normally developed. (B) The same situation experimentally induced by ablation of the left eye and a regenerate antenna-​like structure. (C) A similar case in the stomatopod Rissoides pallidus. Here, the right eye is replaced by a flagellate antenna-​like structure. (D) A right thoracopod situated on the left side of the sixth pleonic segment of a female Carcinus maenas. (E) A thoracopod replacing the endopod of the left fourth pleopod of Panulirus argus. (F) Antenna-​like structures on a pleopod of Panulirus argus. (G) A right third maxilliped of Cancer pagurus with an endopod resembling a cheliped. (Gʹ) The normally developed left third maxilliped of the left side of the same specimen. (A, G, Gʹ) Modified after Bateson (1894). (B) Modified after Herbst (1899). (C) Modified after Giesbrecht (1910). (D) Modified after Bethe (1896). (E, F) Modified after Fausto-​Filho and Da Costa (1977).

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Developmental Biology and Larval Ecology sperm transfer (Gordon 1963). More examples of this sort are a female of Cancer pagurus exhibiting a somewhat reduced cheliped, forming the exopod of a biramous limb in the normally limbless sixth pleon segment (Young 1933) and a female of Carcinus maenas with a fully developed uniramous right pereopod on the left side of the same segment (Bethe 1896) (Fig. 4.1D). The third group includes limb structures in body parts that normally do not show limbs. Dexter (1954) mentioned a strange case of a misplaced limb structure. He found a lobster with a biramous leg-​like structure on the carapace (i.e., at a place that normally does not even show any outgrowth). The most frequent heteromorphosis, however, is the replacement of a stalked compound eye by a structure resembling the first antenna (Fig. 4.1A–​C). This has been found to occur naturally and experimentally induced in a variety of decapods such as shrimp, spiny lobsters, freshwater crayfish, and brachyuran crabs (Bateson 1894, Hofer 1894, Herbst 1896, Steele 1907, Lissmann and Wolsky 1933, Ravindranath 1978, Nevin and Malecha 1991, Scholtz et al. 2014, Rodrigues et al. 2019, Ventura et al. 2019), but also in the stomatopod Rissoides pallidus (Giesbrecht 1910). With very few exceptions, heteromorphoses display more or less complete limbs and not just single-​limb characters such as specific setae, podomere shapes, or serrations. Moreover, the misplaced parts often correspond to the original ones. For instance, the part of the reduced cheliped that replaces the palp in the third maxilliped of crabs shows the same number of podomeres—​namely, merus, carpus, propodus, and dactylus (Bateson 1894, Lira et al. 2013) (Fig. 4.1G, Gʹ). Furthermore, most examples reveal that in cases of homeotic changes of limbs, endopods replace endopods and not exopods [with the notable exception of the supernumerary claw in the pleon of Cancer pagurus (see Young 1933)]. All this indicates the modular nature of heteromorphoses and homoeoses. Fission of Limbs By far, most reports on crustacean malformations deal with multiplications of certain structures of crustacean appendages (Figs. 4.3–​4.6). The multiplied structures resemble the original parts (i.e., the curvature, the equipment with tubercles, spines, and the number of podomeres). However, little is known about internal structures of the multiplied limb parts. It seems that muscles and nerves are somewhat reduced. For instance, Przibram and Matula (1913) studied the anatomy and behaviour of the supernumerary branches of a triplicated second antenna of a spiny lobster and showed the absence of a reaction on external stimuli correlated with reduced nerves. Frequently, the affected structures show triplications, and even in many cases, when only duplicate elements are expressed, a threefold contribution of substructures can be recognized. Hence, Przibram (1921), who studied these phenomena intensely, coined the term Bruchdreifachbildung (BDB; triplicate structures after damage) for this class of malformations. The first accounts of BDB date back to Berniz (1671) and Rösel von Rosenhof (1755), who showed malformed claws in the lobster Homarus gammarus and in the noble crayfish Astacus astacus, displaying varieties of multiplications of parts (Fig. 4.3A, B). Since then, innumerable—​ mostly anecdotal—​accounts of these phenomena have been published. It is impossible to gain a comprehensive picture of all cases and patterns because mentions of these malformations often hide in taxonomic treatments of the various crustacean groups and in many smaller local journals that are difficult to track. Nevertheless, a random count based on some of the older literature—​in particular, the detailed accounts of Bateson (1894), Przibram (1921), and some of the more recent publications—​allows for an estimation of the relative frequency of limb part multiplications. With about 200 published cases, by far most of the limb part multiplications in decapods occur in the large chelipeds (Rösel von Rosenhof 1755, Faxon 1881, Le Sénéchal 1888, Bateson 1894, Herrick 1895, Cole 1910, Dawson 1920, Przibram 1921, Abeloos 1932, Pérez 1936, Shuster et al. 1963, Nickerson and Gray 1967, Gray 1968, Shelton et al. 1981, Nakatani et al. 1997, Feldman 2003, Lira et  al. 2003, Pinheiro and De Toledo 2010, Martínez and Rudolph 2010, Ramírez-​Rodríguez and Félix-​Pico 2010, Matsubara 2011, Schweigert et al. 2013, this chapter) (Figs. 4.3 and 4.4). About a dozen reports deal with the other pereopods (Faxon 1881, Bateson 1894, Andrews 1904, Przibram 1921, Zalpeter 1927, Nickerson and Gray 1967, Lawler and Van Engel 1973; Fausto-​Filho and Da



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Fig. 4.2. Experimentally produced heteromorphoses by artificially misexpression of the ubx gene in the amphipod Parhyale hawaiensis. (A) A thoracopod replacing the right first antenna (highlighted in blue). (B) Transformation of the segments of the second maxilla and the maxilliped to anterior pereopod segments including coxal plates and the transformation of anterior walking legs to a more posterior morphology (highlighted in blue). Modified after Pavlopoulos et al. (2009), with permission from PNAS. See color version of this figure in the centerfold.

Costa 1977, Carmona-​Suarez 1990, this chapter) (Figs. 4.5A, B and 4.6B, C). There is only one case of triplication in a pleopod (Pérez 1926) (Fig. 4.6D). In addition, four descriptions exist of aberrant second antennae of spiny lobsters (Bateson 1894, Przibram and Matula 1913), crayfish (Astacus astacus) (Bateson 1894), and shrimp (this chapter) (Figs 4.5C and 4.6A, E). The experimental ablation of eyes in the shrimp Macrobrachium acanthurus resulted in cases of regenerated antenna-like structures that sometimes showed duplications and triplications (Rodrigues et al. 2019). This is an interesting case of a combination of heteromorphosis and BDB. Most of the reports deal with large and commercially important decapod species. For instance, there are about 70 cases of Homarus americanus, 24 of H. gammarus, 23 of Cancer pagurus, 18 of Astacus astacus, 11 of Carcinus maenas, 10 of Paralithodes camtschaticus, and a similar number of Callinectes sapidus. Furthermore, mostly single examples were found in a number of brachyurans, other astacids, and achelates. Apart from Paralithodes camtschaticus, BDB have been rarely reported in Anomala (Bateson 1894, Pérez 1926, Schweigert et al. 2013). In addition, there are some observations of multiplied limb parts in other crustaceans. For instance, the barnacle Semibalanus balanoides sometimes shows cirri with more than two rami (Stubbings 1975). Lankester (1881) reported an additional small epipodite in the 40th limb of the notostracan Triops cancriformis. Chopra (1934) and Serène (1950) described uropods with duplications of spines in the stomatopods Oratosquillina interrupta and Gonodactylus chiragra. Needham (1950) observed a case of a triplication of the eighth thoracopod of Asellus aquaticus (Fig. 4.6G). Finally, two specimens of the amphipods Parajassa pelagica and Jassa falcata displaying bifurcated dactyli of gnathopods were found (Vader 1968, Moore 1973) (Fig. 4.6F).

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The multiplications concern a number of podomeres of a leg (cheliped or another pereopod) (Faxon 1881, Bateson 1894, Dawson 1920, Nickerson and Gray 1967) (Figs. 4.4H–​K , 4.5A, and 4.6C, G), single podomeres (Figs. 4.4C, F and 4.6B, D, F), or parts thereof (Faxon 1881, Bateson 1894, Przibram 1921) (Figs. 4.4A, B, D, E, G; 4.5B, C; and 4.6A, E). The supernumerary parts are always of the same size. Furthermore, in all cases, the multiplications concern distal limb elements, all originating from an undivided basal part. There is no example in which the branching occurs in proximal podomeres with an undivided distal part of the limb. The most comprehensive multiplication comprises all podomeres from the basis on, as in the case of a walking leg of the spiny lobster Palinurus vulgaris (Bateson 1894) and of the eighth thoracopod of the isopod Asellus aquaticus (Needham 1950)  (Fig. 4.6G), and from the ischium on a cheliped of Paralithodes camtschaticus (Nickerson and Gray 1967) (Fig. 4.4K). Dawson (1920) described a bizarre example of a multiplication of several podomeres in the lobster Homarus americanus (Fig. 4.4J). In this animal, three slightly smaller claws replace the primary right crusher claw. These originate as two branches from the merus; the outer branch forms two mirror-​image crushers with fused carpus and propodus, and the inner branch is a single normally developed nipping claw. However, multiplications comprising several podomeres are quite rare. Perhaps these complex structures reduce the general fitness of the animal and lead to fatalities during the molting process. Hence, with about 180 observed cases, the vast majority concerns the branching of the terminal parts of chelipeds, such as the dactylus and the propodus (Figs. 4.3 and 4.4). There are duplications and multiplications of the dactylus in several directions, including the dorsoventral plane (see Le Sénéchal 1888, Bateson 1894, Shelton et  al. 1981)  (Figs. 4.3B, F, I  and 4.4C, F, I). Similarly, the fixed finger of the chelipeds shows frequent duplications, triplications, or even higher numbers, although this structure is just a process of the propodus and not a proper podomere ( 4.3A, C-H, J and 4.4A, B, E, G) (e.g., Lira et al. 2003). An extreme case of a claw deformation was seen in a specimen of Cancer pagurus, in which the dactylus and the fixed finger show triplications (Le Sénéchal 1888) (Fig. 4.4I). Schweigert et  al. (2013) depicted a claw of the fossil hermit crab Schobertella simonsenetlangi from the Early Jurassic that displays a triplication of the pollex (Fig. 4.3C). This so-​far oldest occurrence reveals that similar mechanisms of regeneration must have been active 250 million years ago. Despite the different parts involved in multiplication, several common principles are shared by most examples found in the literature (Figs. 4.3–4.6; see also Bateson 1894, Shelton et al. 1981): (1) the branching always relates to distal parts that originate from a common stem; (2) the additional branches show the same distal elements as the original structure beginning at the bifurcation; Fig. 4.3. Bruchdreifachbildung in decapod chelipeds. (A)  The possibly oldest account (1671) of this phenomenon showing a right lobster claw (crusher) with duplication of the dactylus and triplication of fixed finger of the propodus. (B) Astacus astacus claw with triple dactylus. (C) The oldest fossil example of a Bruchdreifachbildung. The chela of the Jurassic hermit crab Schobertella simonsenetlangi showing a triple fixed finger. (D) A noble crayfish Astacus astacus with an outer duplication of the right fixed finger. (E) Another specimen of Astacus astacus with a triple fixed finger with the additional elements facing inward. (F) A complicated pattern of horizontal duplications of the fixed finger and the dactylus of the right claw of Astacus astacus. (G) Triple fixed finger of the right claw of the shore crab Carcinus maenas. This image clearly demonstrates the orientation of the multiplied elements. The two outer parts are in the correct orientation; the middle part shows a mirror image (see text). (H) Right claw of a lobster with a horizontal triple fixed finger. As with vertical multiplications, the outer elements are in the right orientation and the middle one is a mirror image. (I) Claw of Cancer pagurus with horizontal duplication of the dactylus. ( J) Lobster claw with the same pattern as the fossil hermit crab in (C). (A) Modified after Berniz (1671). (B) Modified after Rösel von Rosenhof (1755). (C) Photo courtesy of Günther Schweigert. (D–​F) Photos courtesy of Gerhard Scholtz, with two specimen of the Museum für Naturkunde Berlin and one specimen of the Naturalienkabinett Waldenburg. (F) Photos courtesy of Markus Frederich. (H–​J) Photos courtesy of Michel Le Quément from the collection of Charly Le Coadou. See color version of this figure in the centerfold.

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Fig. 4.4. Patterns of Bruchdreifachbildung in decapod chelipeds. (A, B) Right and left claws of the crab Necora puber demonstrating the term Bruchdreifachbildung. The multiplication concerns the fixed finger. (A) A triplication with the general pattern of outer parts in the right orientation whereas the middle part is a mirror image. (B) In this case, the additional structure is a composite of the two separate additional parts seen in (A). Accordingly, the additional structure shows ventral sides (i.e., two elements in addition to the original structure are combined). (C) Cancer pagurus with triplication of the dactylus above the propodus/​dactylus joint. (D) Crab claw with triplication of the dactylus in front of the propodus/​dactylus joint. (E) Triple fixed finger of Nephrops norvegicus with misplacement of the original structure, at first sight resembling the pattern in (D). (F) Horizontal triplication of the dactylus in Cancer pagurus. (G) Horizontal duplication of a fixed finger in the crab Necora puber. (H) Triplication of lobster claws (Homarus americanus). The merus shows two ends: one carpus forms two branches, each bearing a claw; the other carpus is a single structure with one claw. (I) Complex pattern (Cancer pagurus) with a combination of horizontal and vertical multiplication of the dactylus and the fixed finger (each triplicate). ( J) Another lobster with an incomplete triplication of the claws. In this case, the pattern deviates from the general principles of BDB, because the handedness of fused claws should be in the opposite direction. (K) Complex triplication of the cheliped of Paralithodes camtschaticus (see text for explanation). (A, B, I) Modified after Le Sénéchal (1888). (C) Modified after Pérez (1936). (D) Modified after Cole (1910). (E, F) Modified after Shelton et al. (1981). (G) Modified after Abeloos (1932). (H) Modified after Emmel (1907). ( J) Modified after Dawson (1920). (K) Modified after Nickerson and Gray (1967).



Patterns of Malformation in Crustaceans (A)

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Fig. 4.5. Patterns of Bruchdreifachbildung in appendages other than chelipeds. (A) Birgus latro with triplicate second left pereopod. (B) Additional fixed finger in the third pereopod of the crayfish Orconectes limosus. (C) Duplication in the antennal scale (scaphocerite) of the freshwater shrimp Paratya curvirostris. (A) Photo courtesy of Jakob Krieger. (B) Photo courtesy of Gerhard Scholtz. (C) Photo courtesy of Stephen Moore. See color version of this figure in the centerfold.

(3)  mostly, the multiplied parts lie in one plane, either the anteroposterior or the dorsoventral; (4) in cases of duplications, the additional branch shows a mirror image orientation to the original branch; (5) in triplications, the middle branch is mirror image oriented and the two outer branches ahow a normal orientation; (6) depending on the orientation of the multiplication, either the ventral and dorsal or the anterior and posterior sides of the additional parts correspond to those of the original structure; and (7) in few cases, these two planes are mixed (i.e., the additional parts undergo a bifurcation themselves, with an orientation that stands at a right angle to the original part). These rules hold true for the cases of multiplication of several podomeres. However, if relatively short structures such as scaphocerites or the fingers of claws are multiplied a slightly different pattern shows up. (1) bifurcation in the anteroposterior plane does not necessarily lead to a proper mirror image, but produces a composite of two structures; (2) in cases of true triplications, these are separated and show the typical pattern with the middle branch forming a mirror image; (3) the branches originating from the point of bifurcation or multiplication do not necessarily show the same number of distal elements. In addition to these multiplications, the additional parts are frequently distorted and/​or rotated, which leads to an even more complicated appearance. Apparently, limb part multiplications do not necessarily hamper very much the individuals bearing them. Many of the specimens described seem to have lived quite a while with these malformations. The observation by Shuster et al. (1963) of a juvenile female of the blue crab that molted three times with a left cheliped showing a triplication of the fixed finger supports this view. Fusion of Limb Parts The number of reported cases concerning the fusion or incomplete separation of limb parts is very low. The most conspicuous examples are thoracic legs with fused endo-​and exopods found in some specimens of the copepod Apocyclops ramkhamhaengi (Chullasorn et al. 2008) (Fig. 4.6H). One can discuss whether this pattern is the result of a proper fusion or, even more likely, the suppression of the separation of the two limb branches during development. In any case, the resulting structure forms a broad paddle showing the setation of the inner margin of the endopod and the outer margin of the exopod (Fig. 4.6H) (Chullasorn et al. 2008).

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Fig. 4.6. Patterns of Bruchdreifachbildung in appendages other than chelipeds (A–​G). Fusion of limbs (H) and multiplication and fusion of other body structures (I–​K). (A) A triple second antenna of Panulirus argus. (B) A triple dactyl of the third left pereopod of the crayfish Cambarus bartonii. (C)  A  bifurcate merus in combination with a bifurcate dactylus in Paralithodes camtschaticus. (D) A triplication of the terminal segment of the first pleopod of the porcellanid crab Pisidia longicornis. (E) Triple tip of the antennal scale (scaphocerite) of the freshwater crayfish Astacus astacus. (F) Duplication of the dactylus of the gnathopods in the amphipod Jassa falcata. (G) Triplication of the eighth thoracopod of the isopod Asellus aquaticus. (H) Thoracopod of the copepod Apocyclops ramkhamhaengi with a fused endo-​and exopod on the left side. The right side is normal. (I) A shrimp (Athanas nitescens) with a bifurcated rostrum. ( J) Multiplication of the lateral carapace spine in the crab Callinectes sapidus. (K) Fused rostral spines of the spiny lobster Panulirus laevicauda. (A, K) Modified after Fausto-​Filho and Costa (1977). (B) Modified after Andrews (1904). (C) Modified after Nickerson and Gray (1967). (D) Modified after Pérez (1926). (E) Modified after Bateson (1894). (F) Modified after Moore (1973). (G) Modified after Needham (1950). (H) Modified after Chullasorn et al. (2008). (I) Modified after Ashelby and Lavesque (2011). ( J) Modified after Faxon (1881).

OTHER APPENDAGES AND BODY OUTGROWTHS Multiplications and fusion of axes do not only occur in proper limbs. This phenomenon also concerns other appendages or outgrowths of the trunk, such as pointed terminal regions or lateral spines of the carapace. There are a few reports on bifurcations of terminal body structures. A  number of authors described bifurcated rostra or rostral spines in dendrobranchiate (Penaeus californiensis) and



Patterns of Malformation in Crustaceans

caridean shrimps (Athanas nitescens, , Palaemon longirostris, and P. serratus), a lobster (Homarus americanus), and two spiny lobsters (Panulirus argus and P. laevicauda) (Herrick 1895, Fauvel 1900, Fausto-​Filho and Da Costa 1977, Aguirre and Hendrickx 2005, De Grave and Mentlak 2008, Ashelby and Lavesque 2011) (Fig. 4.6I). Likewise, bifurcated telsons and telson spines occur in decapods—​ namely, in specimens of the dendrobranchiates Penaeus vannamei and Parapenaeopsis stylifera, the carideans Leptocarpus potamiscus and Nematopalaemon tenuipes, and the stomatopod Pseudosquilla ciliata (Serène 1950, Dutt and Ravindranath 1974, Aguirre and Hendrickx 2005). These bifurcations follow a pattern similar to that of bifurcated limbs (i.e., the two branches seem to form mirror images to each other). As in biramous limbs, fusion of the paired rostral spines of the spiny lobster Panulirus laevicauda has been observed (Fausto-​Filho and Da Costa 1977) (Fig. 4.6K). Comparable to the situation in the rostrum and the telson, spines at the lateral carapace margins show bifurcations or even trifurcations in the brachyurans Portunus pelagicus and Callinectes sapidus (Faxon 1881, Rasheed et  al. 2014)  (Fig. 4.6J). The figures in these publications indicate that a trifurcated spine at the carapace margin shows the characteristic symmetry situation found in limb trifurcations. Hoch and Yuen (2009) described a specimen of the barnacle Semibalanus balanoides with two penises, which could be the product of a split during early differentiation.

TRUNK Conjoined Twins The most fundamental malformations in crustaceans are the conjoined twins (Figs. 4.7 and 4.8). These are characterized by a partial or complete duplication of the anteroposterior body axis and show various degrees of fusion. There are currently 17 publications reporting more than 20 instances of conjoined twins in 15 species of crustaceans. As is true for all crustacean malformations, most reported cases concern decapods—​in particular, the European and American lobster (Homarus gammarus and H. americanus) (Brightwell 1835, Ryder 1886, Herrick 1895, Harzsch et al. 2000) as well as various crayfish species (Astacus astacus, Cambarus longulus, Cherax destructor, Pacifastacus leniusculus, Procambarus fallax f.  virginalis, and Virilastacus rucapihuelensis) (Reichert 1842, Zipf 1956, Harlioğlu 2002, Alwes and Scholtz 2006, Rudolph and Martinez 2008, Scholtz 2014) (Figs. 4.7B–​D and 4.8D–​F). Furthermore, three cases have been described in three representatives of the Anomala-namely, Aegla abtao ( Jara and Palacios 2001), Aegla uruguayana (Diawol et al. 2019), and Lithodes aequispinus (this chapter) (Fig. 4.7E), a brachyuran crab (Amarinus lacustris) (Scholtz et al. 2014) (Fig. 4.7A), and the shrimp Rimapenaeus similis (Williams 1988) (Fig. 4.8B). By contrast, there are only a few reported examples of conjoined twins in non-decapod crustaceans: one finding by Chatton (1909) concerning four specimens of the copepod Ophioseides cardiocephalus (Fig. 4.8A) and two reports of duplications of the externa in the rhizocephalan Cirripedia Sacculina carcini and Loxothylacus panopaei (Pérez and Basse 1928, Reinhard 1954) (Fig. 4.8C). With more than 10 examples, most crustacean conjoined twins were observed in early larval and early juvenile stages (Brightwell 1835, Ryder 1886, Herrick 1895, Chatton 1909, Zipf 1956, Jara and Palacios 2001, Harlioğlu 2002, Rudolph and Martinez 2008) (Fig. 4.8A, D, E). Ten double embryos have been reported (Reichert 1842, Herrick 1895, Harzsch et  al. 2000, Alwes and Scholtz 2006, Scholtz 2014, Diawol et al. 2019, this chapter) (Figs. 4.7B–​E and 4.8F). Only four cases concern adult animals (Pérez and Basse 1928, Reinhard 1954, Williams 1988, Scholtz et al. 2014) (Figs. 4.7A and 4.8B, C). The relatively rare observation of conjoined twins in late larval stages or adult crustaceans is explainable by the low probability of survival of free-​living conjoined twins. The complex surface structures make the regular molting processes, which are problematic anyway, even more difficult,

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Fig. 4.7. Conjoined twins. (A)  A  specimen of the freshwater crab Amarinus lacustris with three eyes and a dorsal antenna-​like structure (arrows). This has been interpreted as anterior duplication (duplicitas anterior) of the embryo, with fusion of the median parts such as the eyes and first antennae. (B, C) Two examples of embryonic conjoined twins (duplicitas completa) of the crayfish Cherax destructor (lateral views). (B) Two equal-​size embryos (arrows point to the eyes) are conjoined dorsally, sharing the yolk (y). The embryonic caudal papillae (cp) mark the posterior ends. (C) Two unequal-​size embryos (arrows point to the eyes) are conjoined dorsally, sharing the yolk (y). The left embryo is complete, including the posterior caudal papilla (cp). The right one is smaller and incomplete, with a body rudiment (br) lacking posterior structures. (D) Germ bands of conjoined embryos (duplicitas posterior) of the marbled crayfish Procambarus f. virginalis stained with a fluorescent nuclear marker (ventral view). The left (lle, left embryo’s left eye; lla1, left embryo’s left first antenna; llr, left embryo’s labrum; lcp, left embryo’s caudal papilla) and right (rre, right embryo’s right eye; rra1, right embryo’s



Patterns of Malformation in Crustaceans

and locomotion as well as feeding are most certainly impaired too. Conjoined embryonic and larval stages, on the other hand, are seldom found because they are small and difficult to access. Patterns of Conjoined Twins Apart from other patterns observed especially in mammals, including humans (e.g., Spencer 2000, Kaufman 2004), but also in arthropods (e.g., Patten 1896, Brauer 1917), scientists traditionally distinguish between conjoined twins with double anterior structures (such as the head), which are referred to as duplicitas anterior (DA); or twins with double posterior body regions, referred to as duplicitas posterior (DP). By far the most frequently observed pattern in crustaceans is the DP version (Brightwell 1835, Ryder 1886, Herrick 1895, Harzsch et al. 2000, Harlioğlu 2002, Alwes and Scholtz 2006) (Figs. 4.7D and 4.8E, F). This applies to crayfish and lobsters. Only the four metanauplius larvae of the copepod mentioned earlier (Chatton 1909), an adult shrimp (Williams 1988), an adult crab (Scholtz et  al. 2014), and one juvenile crayfish specimen (Rudolph and Martinez 2008) display a DA (Figs. 4.7A and 4.8A, B, C). The shrimp Rimapenaeus similis is unique, because the front parts of the two bodies do not lie next to but on top of each other and thus form a pattern, which is hard to explain. This is likely the result of an abortive molting process than a case of conjoined twins (Williams 1988) (Fig. 4.8B). With two rostra, three eyes, and an additional dorsal antenna-​like structure, the crab Amarinus lacustris shows a pattern that is also difficult to explain and thus only indirectly inferred to as DA (see Scholtz et al. 2014) (Fig. 4.7A). The double structures found in the externa of the two Rhizocephala species are not easy to interpret either, because the extreme modification of the body organization in these parasitic crustaceans conceals the relation of the body axes to each other. In line with the way that most scientists interpret the orientation of the externa, both reported cases represent a DA (Pérez and Basse 1928, Reinhard 1954) (Fig. 4.8C). The situation is a bit different in the cases of dorsal adhesion of the cephalothorax region in newly hatched lobsters, crayfish, and Aegla abtao (Ryder 1886, Zipf 1956, Jara and Palacios 2001, Rudolph and Martinez 2008). This pattern is called duplicitas completa (DC) (Fig. 4.8D). This situation relates to the double embryos, each of which has its own complete anteroposterior body axis, as described by Reichert (1842), Herrick (1895), Scholtz (2014, this chapter), and Diawol (2019) (Fig. 4.7B, C, E). The pattern of dorsally conjoined juvenile stages forms from two separate ventral germ band anlagen, the backs of which fuse while they grow around the yolk (see also Herrick 1895). The two embryos can be of more or less the same size or differently developed (see Scholtz 2014) (Fig. 4.7B, C). In crayfish embryos, as originally in all malacostracans, the posterior thorax region and the pleon are formed by a teloblastic growth zone in a ventrally folded caudal papilla (Scholtz 2000, Alwes and Scholtz 2006). Due to this ventral folding, the posterior thorax and the pleon of conjoined twins are not fused at hatching, even in cases of a DC (see Fig. 4.8D). By contrast, one of the two juvenile DC specimens of the crayfish Virilastacus rucapihuelensis described by Rudolph and Martinez (2008) shows a complete fusion along the body axis, including the entire pleon. This pattern suggests that the two early germ bands must have shared one growth zone. Hence, during early embryogenesis the conjoined twins formed a DA. Only with advanced development, the two anterior body regions fused, resulting in an extreme case of a DC.

right first antenna; rlr, right embryo’s labrum; rcp, right embryo’s caudal papilla) germ bands share a fused eye (fe) and the fused first antenna (fa). All posterior segments are normally developed. (E) Conjoined twins (presumably duplicitas completa) of Lithodes aequispinus (anterior perspective). The two embryos are marked by arrows. (A) Modified after Scholtz et al. (2014), with permission from Elsevier. (B, C) Photos courtesy of Gerhard Scholtz. (D) Photo courtesy of Frederike Alwes and Gerhard Scholtz. (E) Photo courtesy of Bradley G. Stevens. See color version of this figure in the centerfold.

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Fig. 4.8. Conjoined twins. (A) Duplicitas anterior of a metanauplius of the copepod Ophioseides cardiocephalus (ventral view). Two anterior naupliar regions (left naupliar region l, right naupliar region r; they are marked by dotted lines) each equipped with an unpaired nauplius eye (lne, eye of the left naupliar region; rne, eye of the right naupliar region) and paired first and second antennae (e.g., lla1, left first antenna of the left naupliar region; rla1, right first antenna of the left naupliar region; lla2, left second antenna of the left naupliar region; rla2, right second antenna of the left naupliar region). The right mandible of the left and the left mandible of the right naupliar region are conjoined (fmd). The counterparts are normal (e.g., llmd, left mandible of the left naupliar region). The first maxilla is the first normal paired limb. (B) Duplicitas anterior of an adult decapod shrimp Rimapenaeus similis (lateral view). The vertical arrangement of the duplicated cephalothorax region is unusual (see text). (C) Duplicitas anterior of the externa of Sacculina carcini (see text) (st, stalk that connects the externa with the interna in the crab host’s interior). (D) Duplicitas completa of the early postembryonic stage of the South American freshwater anomalan decapod Aegla abtao (lateral view). The dorsal cephalothorax regions of the two individuals are conjoined. This pattern develops from double embryos (as shown in Fig. 4.7B). (E, F) Two cases of a Duplicitas posterior in the American lobster Homarus americanus showing different degrees of conjoined anterior parts (dorsal view). (E) Zoea larva with one median eyespot (e), conjoined first antennae (a1), and partly conjoined cephalothorax region with double pleon. (F) Central nervous system of an embryo. Normal-​appearing anterior brain region with paired anlagen of the eyes (e) and the brain parts protocerebrum



Patterns of Malformation in Crustaceans

Structures that have grown together are frequently observable in DA as well as in DP patterns. Of course, this applies in particular to those parts situated in the region of transition between the single and the double body axes. An example of this is the thicker than normal mandible in the DA median region in the metanauplius of Ophioseides cardiocephalus (Chatton 1909), which obviously consists of parts of the mandibles of both head anlagen (Fig. 4.8A). Another instance includes the conjoined anterior brain parts and the foregut in the DP of Homarus americanus (Harzsch et al. 2000) (Fig. 4.8F). It is furthermore conspicuous that, in many cases of DA and DP, the region that does not show duplication is also malformed and shows various degrees of adhesion. For example, Ryder (1886) and Herrick (1895) described heads with a regular set of paired eyes and antennae for various DP in lobster larvae; however, they also found specimens with only one median eye spot and unpaired appendages (Fig. 4.8E). Alwes and Scholtz (2006) reported a Marmorkrebs DP, which shows the anlagen for a median third eye and a first antenna formed by the corresponding parts of both heads (Fig. 4.7D). The DA metanauplius described by Chatton (1909), also has a wider posterior body region compared to a normal larva (Fig. 4.8A). Strictly speaking, this means that a DA or DP is not necessarily and not always merely a separation of a part of the longitudinal body axis, but rather a partial overlapping of two body axes, resulting in various degrees of adhesion, depending on the angle at which they are situated in relation to each other. In cases when the germ anlagen are parallel to each other, the axes do not initially overlap and thus no adhesions occur. It is not until a later stage that the two embryos inevitably fuse because they are growing on a single, shared yolk mass. Irregular Segmentation In some crustacean specimens, there is a mismatch between the right and left body halves concerning segmental structures (Fig. 4.9). A male specimen of Proasellus cavaticus shows an eighth thoracic segment with a left side that is normal whereas the right half is very small and lacks its leg and the penis (Henry 1966). Likewise, a female king crab Paralithodes camtschaticus reveals a reduced fourth left sternite and an absent fourth left leg (Stevens and Munk 1991). The malformed Ucides cordatus has only four pairs of walking legs on the right body side, whereas the left side is equipped with the regular number of five legs (Araújo and Calado 2012). Rasheed et al. (2014) reported a pleon with a small additional pleomere situated on the left body half between the fifth and sixth pleonic segments. Manning (1962) described a somewhat different case of a female Squilla bigelowi in which the sixth and seventh thoracic segments are normally expressed on the right side with thoracopods 6 and 7 (Fig. 4.9A). However, from the midline to the left side, there is only one segment without any segmental boundary and only one leg. Because the lateral process of this half segment resembles that of the right seventh segment and the gonopore, normally found in the sixth segment, and is present, Manning (1962) concluded that it is a fusion of the sixth and seventh thoracic segments. Linder (1952) and Šaganović et al. (2019) reported several comparable cases of bilateral segmental mismatch in the abdomen of notostracans. (pc), deutocerebrum (dc), and tritocerebrum (tc). Alternatively, this could represent a combination of right head parts (r) of a right embryonic anlage and the left head parts (l) of a left embryonic anlage. In the area of the bifurcation, a conjoined anlage of the protocerebrum and clearly separate anlagen of the deuto-​and tritocerebrum are present. These are the right parts (r) of the left embryo and the left parts (l) of the right embryonic anlage. Beginning with the deutocerebrum, all elements of the central nervous system are complete and paired. mdg, mandible ganglion; oe, paired esophagus, leading to one mouth opening. (A) Modified after Chatton (1909). (B) Modified after Williams (1988). (C) Modified after Pérez and Basse (1928). (D) Modified after Jara and Palacios (2001). (E) Modified after Herrick (1895). (F). Modified after Harzsch et al. (2000), with permission from Elsevier.

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Fig. 4.9. Segmental irregularities. (A) The posterior thorax of the stomatopod Squilla bigelowi with different segment numbers in the right and left body halves (dorsal view). On the right side there are normally developed sixth and seventh thoracic segments. On the left side there is only one segment. (B, C) Spiral segments (helicomery). (B)  Posterior body region of the isopod Porcellio scaber (dorsal view). The seventh and eighth thoracic segments are incomplete, forming a spiral. This begins in the seventh thoracic segment (thick arrow) and ends in the eighth thoracic segment (asterisk). A second spiral begins in the first (not visible and perhaps absent) or second pleonic segment (thin arrow) and ends in the third pleonic segment (asterisk). (C) Posterior body region of the notostracan Lepidurus lynchi (ventral view). The spiral begins in the leg-​bearing region (arrow) and forms five loops before it ends in the posterior-​most segment (asterisk). (A) Modified after Manning (1962). (B) Modified after Keilbach (1958). (C) Modified after Linder (1952).

A further type of malformation closely related to these examples is the phenomenon of spiral segments or helicomery (Morgan 1895). In these cases, segmental rings are not arranged one after the other along the longitudinal body axis, but tergites, pleurites, and sternites show a screw-​like arrangement (Morgan 1895). Among crustaceans, this complicated pattern has been found to occur in notostracans (Linder 1952, Longhurst 1958, Šaganović et al. 2019) (Fig. 4.9C), in an isopod species (Keilbach 1958) (Fig. 4.9B), and in two pentastomid species (Spencer 1892, Heymons 1931). These spirals begin as a half segment form one or more twists around the body and end again in a hemisegment. The pentastomid specimens show a corresponding pattern but, in this case, it occurs in the secondary annuli characteristic for this parasitic group and not in the proper segments (Spencer 1892, Heymons 1931).

EXPERIMENTAL APPROACHES, INTERPRETATION, AND EXPLANATIONS Putting Limbs in the Wrong Place Grafting Heteromorphoses showing similar patterns as the naturally occurring ones have been produced experimentally using various approaches. For instance, ectopic chelae were induced experimentally in a crab by transplantation of cheliped tissue to the leg stump of autotomized pereopods or to eye sockets (Kao and Chang 1996, 1997). Leg grafting in crayfish sometimes led to the differentiation of an exopod in normally uniramous pereopods (Mittenthal 1980). Extirpation experiments in various decapods and isopods revealed that eye regeneration depends on the amount of ablated nervous tissue (Herbst 1896, 1899, 1901). An ablation of eyes including the optic neuropils leads to the regeneration of an antenna instead of an eye. If only terminal eye regions are removed, an eye regenerates. Yet, a recent eye-ablation study in the crayfish Cherax quadricarinatus showed that the ablation of the entire eye stalk including the optic neuropils did not result in any regeneration.



Patterns of Malformation in Crustaceans

If only the ommatidia and lamina were removed, either an eye or an antenna-like structure were regenerated (Ventura et al. 2019). By contrast, Rodrigues et al. (2019) cut off entire eyestalks of the shrimp Macrobrachium acanthurus and gained different patterns of antenna-like structures, whereas Desai and Achuthankutty (2000) reported a regenerated eye after complete eyestalk ablation in Penaeus monodon. These contradictory results reveal that further studies are required. In any case, several studies indicate that these antennae function similar to normal antennae and project to the olfactory brain parts (Maynard and Cohen 1965, Mellon et al. 1989). These examples suggest that irregular regeneration processes probably cause many of the naturally occurring heteromorphoses after an injury or by disturbance of ontogenetic processes. Gene Expression Recent experiments applying modern molecular techniques in the amphipod Parhyale hawaiensis such as transgenic and recent clustered regularly interspaced short palindromic repeats (CRISPR)/​ CRISPR-​associated system 9 (cut out, put in, and edit identified DNA bits and genes) approaches have shown that a shift in the domain of Hox gene expression leads to homoeotic changes. These concern the transformation of mandibles and maxillae to antennae, of pereopods to maxillipeds or to pleopods and vice versa, or the truncation of head segments (Liubicich et al. 2009, Pavlopoulos et al. 2009, Martin et al. 2016) (Fig. 4.2). These experiments explain the homoeotic transformation of limbs and segments adjacent to the regular boundary of Hox gene expression as in the case of a third maxilliped differentiated as the adjacent cheliped (Fig. 4.1G). However, the transformation of a single distant appendage such as a pereopod or an antenna in the position of a pleopod requires another explanation. Moreover, some of the heteromorphic changes do not indicate (serial) homology between the original structures and those replacing them. This is evident for the additional leg structure on the carapace (Dexter 1954). A crab cheliped is definitely an endopod but occurs as an exopod in the case described by Young (1933), and a right leg is not in a proper position when occurring on the right side (Bethe 1896) (Fig. 4.1D). Likewise, a homology between eyestalks and antennae as suggested by the frequently reported transformations of eyes into first antenna-​ like structures (Fig. 4.1A–​C) is doubtful, to say the least. These instances are more comparable to experiments on the induction of ectopic eyes on wings, legs, and antennae in Drosophila (Halder et al. 1995), which do not imply homology between the wing margin and the head. Atavisms Some authors interpreted heteromorphoses as atavisms showing an ancestral character state within a particular animal taxon (see Giesbrecht 1910, von Buddenbrock 1954, Riedl 1975, Mittenthal 1980, Martin et al. 2016). However, it is evident that this is often not the case. It is difficult to conceive that the malacostracan stem species had an additional pair of first antennae instead of eyes. Likewise, the ancestral decapod certainly did not have second antennae in the pleon, a limb on the lateral carapace, or claws in the third maxilliped. Interestingly, symmetric hermit crabs of the genera Pylocheles and Cheiroplatea have chelate third maxillipeds (Richter and Scholtz 1994), but this is clearly an apomorphy of these groups and no indication for an ancestral character state. Nevertheless, the cases in which legs appear in limbless segments, as in the pleon of brachyurans (Bethe 1896, Young 1933, Gordon 1963), allow for an interpretation as indication of a former presence of limbs. This conclusion is based on the homologous segmental position of the heteromorphic legs compared with those of other decapods such as shrimp and lobsters. However, the atavism relates only to the level of a generalized leg, not the specific expression of a pereopod or gonopod. Furthermore, the idea of an atavistic leg in a limbless brachyuran pleon segment can only be specified because there is good evidence from other decapods and malacostracans that have well-​developed pleopods and uropods.

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Developmental Biology and Larval Ecology Multiplication of Limb Parts Creating Multiplications Most reported cases of BDB concern chelipeds. This is explainable by the prominent role of these appendages for food processing, defense against predators, mating, and combat with conspecifics (Shuster et al. 1963). All this increases the chance for injuries of chelipeds. Likewise, the malformed second antennae in spiny lobsters (Bateson 1894, Przibram and Matula 1913, Fausto-​Filho and Da Costa 1977) find an explanation in the use of these strong appendages in social interactions of these clawless animals (e.g., Lavalli and Herrnkind 2009). This indicates that the main cause for the multiplication of limb structures is an irregular wound healing and regeneration process—​a view that has already been put forward by Rösel von Rosenhof (1755) and was disputed by Bateson (1894). Nevertheless, the grafting and extirpation experiments of Reed (1904), Emmel (1907), Needham (1950), Mittenthal (1981), Kao and Chang (1996), and Nakatani et  al. (1998) support this view. These researchers were able to create BDB artificially in lobster, crayfish, hermit crab, crab, and isopod species. Several authors discussed the possibility of environmental stress by pollutants as being the cause for BDB (Pinheiro and De Toledo 2010). However, in crustaceans there is so far no direct proof for this. Yet, embryos of other arthropods exposed to chemicals or high temperatures showed duplications of limb axes resembling those in crustaceans (e.g., Girton 1981, Napiórkowska et al. 2015, 2016). In any case, the idea that extra limb parts are the result of mutations and thus inherited (see Bateson 1894) is very unlikely. The specific symmetries of BDB led to a number of models implying gradients of morphogens, positional information, boundaries between different cell states, and emergent properties of the circuitry and the spatial arrangment of signalling pathways as explanation for the observed phenomena (Przibram 1909, Abeloos 1932, Shelton et al. 1981, Meinhardt 1983, 2009, Held and Sessions 2019) (Fig. 4.10). In particular, Meinhardt’s Boundary Model shows a good fit with the observed patterns of limb duplications in crustaceans and other arthropods (Fig. 4.10B–Eʹ). According to the Boundary Model, limbs and other lateral branches of the body axis are formed at a boundary where cell populations with at least three different states meet (Meinhardt 1983, 2009) (Fig. 4.10B-Bʺ). Drosophila research has shown that each segment comprises transverse cell populations with an anterior and a posterior fate—​the compartments—​which lie strictly separated but adjacent to each other (Martinez Arias and Lawrence 1985). In addition, the model implies that there is a longitudinal boundary separating dorsal and ventral cells on either lateral side of the embryo (Meinhardt 1983, 2009) (Fig. 4.10B). In the contact zone between anterior and posterior cells and the dorsoventral border, the formation of limb buds is initiated (Fig. 4.10B). Meinhardt’s model has been corroborated twofold: by clonal studies of crustacean segmentation and limb differentiation, and by molecular genetic investigations on Drosophila limb formation. Clonal studies on limb development in crustaceans showed that each leg is composed of cells from two adjacent genealogical units or parasegments (Dohle and Scholtz 1988, Wolff and Scholtz 2008). The early limb buds are formed at a clonal boundary (Hejnol and Scholtz 2004) and in a distinct distance to the midline, which plays a central role for dorsoventral patterning by secreting morphogenetic proteins (Vargas-​ Vila et al. 2010). Drosophila studies revealed that the anterior and posterior cell populations express different segment polarity genes such as wingless and hedgehog/​engrailed, whereas the dorsoventral boundary is marked by the decapentaplegic gene. At the intersection of these genes, the homeobox gene Distal-​less is activated, which initiates the budding of limbs (Campbell and Tomlinson 1995). Meinhardt’s model implies a third segment–​polarity cell state that separates the posterior cells from the anterior cells of the next following segment (Fig. 4.10B). Otherwise, legs with mirror-​image orientation would form at every posterior–​anterior boundary. Again, Meinhardt’s implication finds support from gene expression data. In each normally developing segment anlagen, there is always an anterior region of cells that neither express engrailed nor wingless (e.g., Damen 2002).

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Fig. 4.10. Models explaining the patterns of limb part multiplications and Bruchdreifachbildung. (A, Aʹ) Hans Przibrams models show patterns as a result of regeneration after specific types of injuries. Ventral and dorsal cell states are indicated by red and white areas. According to Przibram, cells tend to regenerate cells of the state that has been removed by the injury. (A) Model explaining the origin and resulting pattern of duplications along the proximodistal axis of a limb. A vertical injury separating the dorsal and ventral cells leads to regeneration of the absent cell states. In this way, a pattern of two-​limb regenerates forms that shows the same dorsoventral orientation. However, such a pattern seems to occur rarely in nature (if at all), because an injury dividing the tip of a limb into two exactly equal-​ size lateral halves is an unlikely event. Furthermore, the contact of cells with chirally arranged opposite states in the notch between the regenerated branches is likey to induce an additional tip with a mirror image orientation. Duplications of the rostrum might be a possible example for this (although in a left–​right direction, see Fig. 4.6I). (Aʹ) Model explaining Bruchdreifachbildung. In this case, a lateral injury affecting either the dorsal or ventral cells

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Developmental Biology and Larval Ecology Fig. 4.10. continued. of the limb and a loss of the tip causes regeneration. As in the previous model, cells of the missing opposite state are regenerated, leading to the specific triplicate pattern, with the two outer regenerates showing the right orientation and the middle structure having a mirror-​image orientation. This is independent of the degree of fusion of the regenerates (see Figs. 4.3–​4.6). (B–​Eʹ) The Boundary Model of Hans Meinhardt explaining the formation of limbs and of multiplication of limb axes based on the boundaries between different cell states. (B–​Bʺ) Formation of arthropod limbs. (B) Schematic representation of segmentation and limb formation. Different colors mark the cell states. Anterior–​dorsal (AD), blue; anterior–​ventral (AV), green; posterior (P), red; separating cell state (S), gray; forming limb, yellow; orientation of the limb, circle with turquoise arrowhead. The limbs form in the area where at least three different cell states meet (AV, AD, P). The topographical relation between ventral and dorsal cells determines the limb orientation as right or left. The gray separating cell state is a requirement to avoid limb formation between the posterior cells of a more anterior segment and the anterior cells of the adjacent segment. These limbs would have a reverse orientation. (Bʹ) Computer model with different cell states (ventral, V; dorsal, D; anterior, A; posterior, P) inducing limb bud growth in the third dimension at the point where these cells stage meet. (Bʺ) The morphological outcome: normal leg. (C–​Cʺ) Explanation of Bruchdreifachbildung with partial fusion of the additional limb. (C) Cell state within a segment with a misplaced patch of posterior cells [color code and abbreviations as in (B)]. The misplaced patch of posterior cells induces additional boundaries between three cell states, resulting in additional limb anlagen. These show the typical orientation, with the middle anlage in a mirror orientation (marked by rotating arrows). (Cʹ) Computerized model of this situation. (Cʺ) The morphological outcome: Bruchdreifachbildung with partly fused additional branches. (D, Dʹ) Similar situation as in (C), but the misplaced patch of posterior cells has a different size and shape. (D) The model. (Dʹ) The morphological outcome: two complete legs with mirror image. (E, Eʹ) Similar situation as in (C), but the misplaced patch of posterior cells has a different size, shape, and position. (E) The model. (Eʹ) The morphological outcome: Bruchdreifachbildung as in (C), but with a complete fusion of the additional limbs. (A, Aʹ) Modified after Przibram (1909). (B–​Eʹ) Modified after Meinhardt (1983, 2009), with permission from Elsevier. See color version of this figure in the centerfold.

A perturbation of cells with different specifications caused by heat, chemicals, or injuries could lead to misplacement of patches of regionally specified cells. This creates additional boundaries between different cell stages. For instance, if a patch of posterior cells appears at the anterior margin of a segment anlage, new intersections of anterior–​dorsal, anterior–​ventral, and posterior cell populations occur. Depending on the size and position of the misplaced cell group, different patterns of duplications or triplications with specific handedness result. Yet, the entire multiplications lie on the same plane. For example, a duplication with mirror-​image symmetry, or a partial triplication with the two outer parts showing the same, the central part an opposite handedness are formed (Fig. 4.10C–C’’). The degree of fusion between these additional limb parts depends again on size and position of the misplaced patches with respect to the original limb bud anlage (Fig. 4.10C–E’). Indeed, this model is in good agreement with the observed patterns of most BDB in crustaceans and arthropods in general. However, to apply the boundary model to processes during regeneration in adult crustaceans, one has to assume that the embryonic clonal boundaries are maintained in fully developed legs and/​or that the regeneration process shows similarities to embryonic development. Both are likely but unknown in crustaceans. Furthermore, Meinhardt’s Boundary Model does not cover the frequently occurring duplications of the dorsal/​ventral axis of limbs, which are sometimes combined with anteroposterior duplication events. Because the dorsoventral multiplications show the same handedness relationships as the anteroposterior ones, and because they are also arranged in one plane, a similar mechanism of translocated cell fates can be assumed. However, this hypothesis needs to be elaborated further. Fusion or Suppression of Separation of Limb Rami A comparison of uniramous thoracopods and biramous pleopods of isopods and amphipods reveals that both limb types begin as limb buds that, only later in development, are bifurcated in the case of the biramous limbs. The early buds of both limb types not only show similar shapes and sizes, but also the same number of individually identifiable cells of theses buds express distal-​less, a homeobox gene involved in the early budding of limbs (Hejnol and Scholtz 2004). Furthermore, a



Patterns of Malformation in Crustaceans

clonal analysis of leg formation in the amphipod Cryptorchestia garbinii revealed that the two clones that constitute the exo-​and endopods of the pleopods together form the uniramous thoracic limbs (Wolff and Scholtz 2008). Given that the biramous limb type is ancestral to the uniramous one, this finding suggests that the suppression of the bifurcation of the early limb bud leads to the uniramous condition. One can assume that something similar has happened in the case of the fused exo-​and endopod in the thoracic limb of the copepod Apocyclops ramkhamhaengi (Fig. 4.6H) (Chullasorn et al. 2008). As a result of some impact on the embryonic limb bud, the separation into an endo-​ and exopod was suppressed. Again, experiments in other arthropods suggest this possibility (e.g., Napiórkowska et al. 2015). Alternatively, an injury of the central region of the biramous limb could have caused a fusion along the wound that led to a broad uniramous paddle. Both scenarios may be also applicable to the fused rostral spines of the spiny lobster Panulirus laevicauda (Fig. 4.6K) (Fausto-​Filho and Da Costa 1977). Yet, the ontogenetic formation of theses spines is unknown (i.e., whether they form from a common bud or grow out independently). Conjoined Twins: Fusion or Incomplete Separation? The causes for the formation of conjoined twins are not fully understood. The general two questions are as follows: (1) At what embryonic stage are conjoined twins formed? (2) Are they formed by the fusion of two early embryo anlagen or as a result of incomplete separation of one embryo anlage? This issue is subject to a controversial debate, even for cases observed in humans (e.g., Spencer 2000, Kaufman 2004). A number of different hypotheses have been put forward concerning the causes for their occurrence in crustaceans. Zipf (1956) postulated that dorsally conjoined crayfish form by fusion of two embryos developing within one egg membrane. Ryder (1886) explained the DP found in the American lobster by the fusion of two embryo anlagen during the course of gastrulation. According to his hypothesis, the angle of the embryos’ longitudinal axes to each other determines the degree of fusion. However, he does not explain how and why the center of gastrulation is duplicated inside the egg before the fusion takes place. Based on his own observations of double embryos in American lobsters, which were all completely separated, Herrick (1895) disagreed with Ryder’s opinion. He suggested that fusion does not take place until the relatively late stage of germ band differentiation. However, Herrick did not address the causes for the duplication of the embryonic anlagen either. Harlioğlu (2002) assumed that the crayfish twin develops during the early germ band stage, and the development of the carapace then prevents the development of two separate embryos, resulting in a DP. The pattern of the Marmorkrebs double embryos, which conjoin in the anterior region, led Alwes and Scholtz (2006) to conclude that these conjoined twins must have developed not later than in the blastodisk stage. For the two cases of conjoined twins observed in Rhizocephala, Pérez and Basse (1928) and Reinhard (1954) suggested that a separation of the early infective stage already led to the development of twins. The fusion of two oocytes or fertilized eggs can almost certainly be ruled out because the yolk content and size of the eggs of conjoined twins equals that of single, regular embryos. None of the authors mention any abnormalities concerning these parameters in double embryos. Herrick (1895) expressly mentioned that, apart from being two embryos, a lobster double embryo he observed was otherwise normally developed. The eggs of the double embryos of Cherax destructor described by Scholtz (2014) do not deviate from other eggs of this species in terms of size or yolk content either. The existence of two complete and fully separated germ bands on one yolk (Reichert 1842, Herrick 1895, Scholtz 2014) suggests a duplication of the early germ anlage. This doubling may—​ induced by internal or external causes—​already be established at the level of axis-​forming gradients in the early zygote, during cleavage, or by division of an originally single germ anlage. Hence, in any case, an initial separation process leading to a duplication of the germ anlage has to be assumed. Two fully separated germ bands do not yet constitute conjoined twins because they only share extra-​embryonic (i.e., nonformative) regions. The two individuals do not fuse to form conjoined twins until later stages of their development.

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Developmental Biology and Larval Ecology Partially conjoined early-​stage embryos, as described by Harzsch et al. (2000) and Alwes and Scholtz (2006) are more difficult to explain. The question here is whether two initially separate germ anlagen fuse in these cases or whether a single anlage is incompletely separated. The occurrence of various degrees of overlapping structures in larvae and other postembryonic stages, and the different angles of the embryos in relation to each other reported by Reichert (1842), Ryder (1886), Herrick (1895), Zipf (1956), Harlioğlu (2002), Harzsch et al. (2000), Jara and Palacios (2001), Alwes and Scholtz (2006), and Rudolph and Martinez (2008) suggest a uniform process of germ anlagen doubling. Accordingly, completely separated double embryos only constitute an extreme case of a continuum of embryos with different distances between them and different positioning angles in relation to each other. Depending on the angle between them, this results either in a DA or in a DP. With regard to the position and the distance between the two embryos, a small or larger fusion will eventually occur. Fusions of separate germ anlagen are the result of partial overlapping and ensuing adhesion of the affected structures. Such overlaps are either the result of the migration of germ anlagen or of the expansion during growth and differentiation of adjacent germ anlagen. However, migration of differentiated germ disks can almost certainly be ruled out, whereas overlapping of parts of germ anlagen, which are separate but close to each other during growth, is quite plausible. In any case, a fusion has to take place in the yolk-​rich eggs of arthropods because the separate germ anlagen are situated on just one yolk mass, even if they are initially completely separated. At the latest, the twins conjoin morphologically with hatching. The early fusions affect ventral and terminal aspects of the developing embryo such as the nervous system, appendages, or the forehead, whereas later fusions affect dorsal parts such as the carapace, tergites, or the heart. The examples in crustaceans demonstrate that the simple alternatives—​separation or fusion—​ as possible causes for conjoined twins are not sufficient to explain the observed patterns and processes. In reality, it has to be assumed that both separation and fusion are involved in the formation of conjoined twins in crustaceans. First, a separation of the germ anlagen has to take place at a very early stage. These individual embryos then fuse to different degrees at different stages of embryonic development, depending on the distance between them and the position of the embryos’ axes in relation to each other. This applies at least to crustaceans with eggs, which are rich in yolk and undergo superficial cleavage. Similar processes seem to apply to other arthropods with a corresponding mode of development, as examples in scorpions (Brauer 1917), Spiders (Oda et al. 2019), xiphosurans (Patten 1896), and myriapods ( Janssen 2013) show. The latter author, however, assumes that the DP in embryos of Glomeris marginata constitutes a division of the growth zone in combination with an unseparated head. Perhaps are indeed different mechanisms, like the formation of two body axes with overlap and a partly separation of the original body axis, involved in the generation of conjoined twins. The outcome of the transplantation experiments of Oda et al. (2019) suggests the first possibility, whereas their laser-ablation experiments speak in favour of the second mode. It remains to be explored how far corresponding processes are found in arthropods eggs that have only little yolk and undergo total cleavage. The examples of conjoined twins in copepods, rhizocephalans, and possibly penaeid decapods (Chatton 1909, Pérez and Basse 1928, Reinhard 1954, Williams 1988), which undergo total cleavage, show that the patterns of conjoined twins are quite similar to those found in arthropods with superficial cleavage. However, descriptions of early stages of ontogeny such as double embryos are completely lacking up to the present day. So far, there are no experimental studies on conjoined twins in crustaceans. Hence, one has to rely on experiments carried out in other arthropods showing similar early development as crustaceans. Arthropod conjoined twins have been artificially produced using a variety of approaches, such as transplantation experiments (Oda et al. 2019), exposing early embryos to heat (Mikulska and Jacuński 1970), centrifugation (Ehn 1962), X-rays (Seitz 1966), laser beams (Oda et al. 2019), or toxic substances (Itow and Sekiguchi 1979, Itow et al. 1998). However, not all these



Patterns of Malformation in Crustaceans

factors are equally important for naturally occurring conjoined twins. For instance, it is highly unlikely that embryos are exposed to the forces occurring during centrifugation. Furthermore, what happens at the cellular level that leads to the malformations during development remains largely unclear. In particular, the research of Itow and colleagues on malformations in xiphosuran embryos sheds some light on the problem (Itow and Sekiguchi 1979, Itow et  al. 1998). The exposition of embryos to heavy metals, and in particular to calcium-​free seawater and to sodium hydrogen carbonate, led to a variety of malformations, including conjoined twins. Itow et al. (1998) explained this with defects in cell migration and cell adhesion, which hamper a proper formation of the germ disk and germ band. Xiphosurans show a largely similar early development as lobsters and crayfish with a yolk rich egg and a relatively small germ disk that forms by cell migration. Hence, similar factors might lead to conjoined twins in crustacean species. Irregular Segmentation Segmental mismatches between the left and the right body halves and helicomery are either the result of regeneration after damage in larvae or adults, or of irregular embryonic development. In the first case, the lack of tissue on one side leads to wound healing that bridges two or more segments, leading to an asymmetric condition. During embryonic development, a mismatch can occur if cells that express segmentation genes are somewhat distorted. This is possible because the early segmental primordia are only a few cells wide (see Scholtz and Dohle 1996, Scholtz 2020). A  number of experiments have induced distortion of embryonic cells using different methods. Again, crustaceans have been somewhat neglected concerning experimental studies dealing with these kinds of trunk malformations. Most investigations are on hexapods and chelicerates. For instance, Balazuc (1955) induced segmental mismatch and helicomery in a mantid by mechanical stimulation (vibration) of the egg case. Spiral segments were observed in larval and adult moths (Tineola bisselliella) after ultraviolet radiation of eggs and embryos (Lüscher 1944). An exposure of opilionid eggs to increased temperatures resulted in some examples of helicomery among other malformation patterns ( Juberthie 1968). In the crustacean Artemia, spiral segments in the abdomen were produced based on an experimental treatment with mycophenolic acid, which has a cytostatic effect (Hernandorena 1993). In Drosophila, null mutants for the morgue gene generated, among other effects, spiral segments in the abdomen of adult flies (Schreader et  al. 2010). Because the morgue protein plays a crucial role in the regulation of cell death, its absence in null mutants affects cell proliferation, which leads to a mismatch of forming segments.

PERSPECTIVES It is obvious that we are far away from a detailed understanding of causes and mechanisms leading to the various patterns of crustacean malformations as described in this short review. A variety of experimental studies on crustacean embryos and adult regeneration—​from the genetic via the cellular to the tissue levels—​is required to gain a new perspective on axis multiplications and deformations. Furthermore, the relation between the environmental stress of all sorts and distinct patterns of malformations should be studied in more detail to allow for a better prediction and analysis of impacts of spills on the crustacean fauna. On the other hand, the bearing of malformations on the understanding of morphology and evolution needs a deeper exploration. It is fascinating to see that morphological structures not only have the potential to repair and regenerate existing parts, but also the potential to produce supernumerary structures. What is more, these supernumerary structures are often well patterned and organized, and are more or less complete.

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ACKNOWLEDGMENTS I thank the late Hans Meinhardt for many discussions concerning deformed limbs. Markus Frederich, Gianna Innocenti, Jakob Krieger, Charly Le Coadou, Michel Le Quément, the late John J. McDermott, the late Stephen Moore, Pierre Noel, Declan Quigley, Eric Rudolph, Günther Schweigert, Bradley G. Stevens, and Judith Weis shared tips, literature, images, and knowledge of crustacean malformations with me. I express my wholehearted thanks to all of them. I am grateful to Oliver Coleman for the loan of specimens from the crustacean collection of the Museum für Naturkunde Berlin.

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Rasheed, S., J. Mustaquim, and S. Khanam. 2014. Some external abnormalities found in edible crabs, Portunus pelagicus and P. sanguinolentus, of Pakistan. Pakistan Journal of Zoology 46:541–​548. Ravindranath, K. 1978. An unusual abnormality in a freshwater shrimp (Decapoda, Atyidae). Crustaceana 35:92–​93. Reed, M.A. 1904. The regeneration of the first leg of the crayfish. Archiv für Entwicklungsmechanik der Organismen 18:307–​316. Reichert, K.B. 1842. Sitzber. S. 43–​44. (1912): 21. Juni 1842. Sitzungsberichte der Gesellschaft Naturforschender Freunde zu Berlin 1839–​1859. Friedländer und Sohn, Berlin, Germany. Reinhard, E.G. 1954. A case of conjoined twins in Loxothylacus (Crustacea, Rhizocephala). Proceedings of the Helminthological Society 21:67–​71. Richter, S., and G. Scholtz. 1994. Morphological evidence for a hermit crab ancestry of lithodids (Crustacea, Decapoda, Anomala, Paguroidea). Zoologischer Anzeiger 233:187–​210. Riedl, R. 1975. Die Ordnung des Lebendigen. Paul Parey, Hamburg, Germany. Rösel von Rosenhof, A.J. 1755. Der monatlich herausgegebenen Insecten-​Belustigung dritter Theil. Nürnberg, Germany. Rodrigues, M.M., L.S. López Greco, and G. Bertini. 2019. First reported occurrence of a heteromorph antennule in the freshwater prawn Macrobrachium acanthurus (Wiegmann, 1836) (Decapoda, Caridea). Crustaceana 92:505–511. Rudolph, E.H., and A.W. Martinez. 2008. Conjoined twins in the burrowing crayfish, Virilastacus rucapihuelensis Rudolph and Crandall, 2005 (Decapoda, Parastacidae). Crustaceana 81:1347–​1355. Rudolph, E.H., and A.C. Verdi. 2010. Intersexuality in the burrowing crayfish, Parastacus pilimanus (von Martens, 1869) (Decapoda, Parastacidae). Crustaceana 83:73–​87. Ryder, J.A. 1886. The monstrosities observed amongst recently hatched lobsters. American Naturalist 20:742–​743. Šaganović, I., S. Makarov, L. Lučić, S. Pavković-Lučić, V. Tomić, and D. Miličić. 2019. Developmental and other body abnormalities in the genus Lepidurus Leach, 1819 (Crustacea: Notostraca) from Serbia. Arthropoda Selecta 28:65–72. Sagi, A., D. Cohen, and Y. Milner. 1990. Effect of androgenic gland ablation on morphotypic differentiation and sexual characteristics of male freshwater prawns, Macrobrachium rosenbergii. General and Comparative Endocrinology 77:15–​22. Scholtz, G. 2000. Evolution of the nauplius stage in malacostracan crustaceans. Journal of Zoological Systematics and Evolutionary Research 38:175–​187. Scholtz, G. 2008. On comparisons and causes in evolutionary developmental biology. Pages 144–​159 In A. Minelli and G. Fusco, editors. Evolving Pathways: Key Themes in Evolutionary Developmental Biology. Cambridge University Press, Cambridge, UK. Scholtz, G. 2014. Verbundene Zwillinge bei Krebsen. Sitzungsberichte der Gesellschaft Naturforschender Freunde zu Berlin (N.F.) 50:173–​188. Scholtz, G. 2020. Segmentation: A Zoological Concept of Seriality. (in press) in A. Chipman, editor. Cellular Processes in Segmentation. CRC Press, Boca Raton, USA. Scholtz, G., and G. Brenneis. 2016. A specimen of Pycnogonum litorale (Arthropoda, Chelicerata, Pycnogonida) with a supernumerary leg is in agreement with the ‘boundary model’ of appendage formation. The Science of Nature 103:13–​13. Scholtz, G., and W. Dohle. 1996. Cell lineage and cell fate in crustacean embryos: a comparative approach. International Journal of Developmental Biology 40:211–​220. Scholtz, G., P.K.L. Ng, and S. Moore. 2014. A crab with three eyes, two rostra, and a dorsal antenna-​like structure. Arthropod Structure and Development 43:163–​173. Schreader, B.A., Y. Wang, S. Carter, J. Grigas, and J.R. Nambu. 2010. Drosophila morgue influences cell numbers and positions in the embryonic nervous system. International Journal of Developmental Biology 54:1425–​1433. Schweigert, G., R. Fraaije, P. Havlik, and A. Nützel. 2013. New Early Jurassic hermit crabs from Germany and France. Journal of Crustacean Biology 33:802–​817. Seba, A. 1735–​64. Locupletissimi rerum naturalium thesauri. Reprint 2001, Taschen Verlag, Köln, Germany. Seitz, K.A. 1966. Normale Entwicklung des Arachniden-​Embryos Cupiennius salei Keyserling und seine Regulationsbefähigung nach Röntgenbestrahlungen. Zoologische Jahrbücher, Anatomie und Ontogenie der Tiere 83:327–​447.

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5  HATCHING

Martin Fritsch, Jørgen Olesen, Ole Sten Møller, and Günther Loose

Abstract Hatching in crustaceans is an active mechanism in which free, mobile individuals are released from the egg envelopes. For the majority of species, this marks the transition from the embryonic phase of the life cycle, which is spatially constrained by the egg, and the free-​living phase. The hatching process of crustaceans has so far not been subject to a detailed comparative treatment across taxa and thus we know little of the diversity of mechanisms, timing in relation to other developmental processes, or evolutionary history. Here we attempt to provide an overview of this diversity throughout the Crustacea. To this end, we treat a particular set of subjects that we consider relevant to the hatching process: the morphology of the involved structures (egg membranes, specialized hatching structures of the hatchling, morphology of the hatchling itself), mechanics of hatchling release, biochemical processes involved in egg shell degradation, maternal and embryonic control and initiation of hatching, as well as the temporal pattern of hatching-​related events. A common feature of the hatching mechanism in the majority of crustacean species is an osmotic swelling of the embryo caused by active water uptake prior to hatching, which builds up pressure against the inside of the envelopes. The remaining features vary according to developmental mode and ecological parameters, but the causality behind many hatching-​related features remains unclear. However, we conclude that the particular life history strategy can have a strong impact on the relative timing of hatching events.

INTRODUCTION Hatching in crustaceans is an active mechanism in which free, mobile (swimming, crawling) larvae or juveniles are released from an inner egg (vitelline) and an outer egg (chorion) membrane after Developmental Biology and Larval Ecology. Edited by Klaus Anger, Steffen Harzsch, and Martin Thiel. © 2020 Oxford University Press. Published 2020 by Oxford University Press.

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Developmental Biology and Larval Ecology embryogenesis. The hatching process of crustaceans has so far not been subject to a detailed comparative treatment across taxa and thus we know little of the diversity of mechanisms, timing in relation to other developmental processes, or its evolutionary constraints. In general, water uptake is required for the release of hatchlings from crustacean eggs. This uptake is the result of an increasing osmotic pressure, which eventually leads to swelling of the inner egg content (both embryo and inner egg membrane) and ends in the rupture of the chorion (outer egg membrane) and hatchling release. In other arthropods (i.e., insects, myriapods, and chelicerates), larvae or juveniles hatch as a result of proteolysis followed by rupturing or splitting of the egg capsule, without water uptake or swelling (exceptions are xiphosurans and pycnogonids that show no proteolysis of the egg membranes) (Roonwal 1944, Bernays 1972, Punzo 2000, Brenneis personal communication 2016). In crustacean life history, hatching marks the transition from the embryonic to a free-​living phase. It is followed by either a larval or juvenile phase. For taxa with internal or protected brood care (e.g., under the carapace valves as in Amphionidacea, Ascothoracida, Cladoceromorpha, Leptostraca, and Peracarida), the free life of the larva or juvenile starts after the release from the brood chamber or pouch, after shedding of the egg membranes (Heegaard 1969, Olesen 2014a,b,c, Olesen and Richter 2013, Høeg et al. 2014). In these cases, the hatching event takes place within the brood chamber or pouch, before the larva or juveniles are finally released. Thus, a late phase of morphogenesis and tissue differentiation can occur in an environment that lacks the spatial constraints of a spheroid egg membrane (e.g., in the embryo-​like cladoceromorphan larvae that develop under the carapace of the mother animal) (Fritsch and Richter 2012, Fritsch et al. 2013). The hatching process comprises several events. Figure 5.1 schematically sums up the progression of events in the hatching process, based on the available literature. First, enzymatic degradation reduces the rigidity of the chorion. The developing embryo and (in most crustaceans) the vitelline membrane starts to expand by water uptake through an increasing osmotic pressure caused by a change of the inner egg osmolarity and membrane permeability. Hence, the tough outer chorion is subjected to mechanical stress from inside the egg. Prehatching larvae or juveniles commonly become physically active near the end of embryogenesis. When the tissues of the hatchling are differentiated and distinct morphological features are apparent, the thus-​far irregular muscular twitching

Egg deposition Developing embryo Differentiation/formation of osmoregulatory organs Regulation/change of egg fluid osmolality Enzymatic degradation of the chorion Water inflow due to osmotic pressure Swelling and growing of the embryo and vitelline membrane Burst/rupture of the chorion Release of embryo within the vitelline membrane Repture of the vitelline membrane Final hatching of locomotory larvae or juveniles

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Fig. 5.1. Hatching process timeline in crustaceans. This scheme exemplifies an overview of the events that might occur during development (embryogenesis) and before the final hatching of crustaceans. Note: Not all events need to be present or passed through until the final hatching of a crustacean larva or juvenile. The order of events can vary within the highly diverse Crustacea, as well as the actual occurrence of each single event. Arrowheads indicate the starting point of an event in the hatching timeline.

Hatching is replaced by regular, controlled contractions and appendages or the entire body begins to move. This facilitates removal of the egg membranes. In species with internal brood care [e.g., in the mysid Americamysis bahia (Wortham-​Neal and Price 2002)], however, parental hatching support is less pronounced, often missing completely. The hatching process is often triggered by environmental factors such as temperature, length of photoperiod, freezing, variation in oxygen levels, or osmotic conditions. In this chapter we review hatching diversity throughout Crustacea with particular focus on developmental timing and mechanistic aspects. Terminology hatching: The hatching process in crustaceans is an active mechanism in which free, mobile hatchlings (larvae or juveniles) are released from an inner egg (vitelline) and an outer egg (chorion) membrane. The hatching process is induced by the embryo, the mother animal, or by both. crustacean egg: The crustacean egg contains a developing embryo and a variable amount of yolk. The embryo and yolk mass are enclosed by two membranes (or shells): the inner chitinous and expandable vitelline membrane, and the outer more inflexible cuticular chorion (e.g., Mawson and Yonge 1938, Marshall and Orr 1954, Davis 1959, Linder 1960, Pandian 1970). In case of species with resting eggs, a third (tertiary) shell can be present (Zaffagnini and Minelli 1970, Belk 1987, Madhupratap et al. 1996, Dumont and Negrea 2002). inner egg (vitelline) membrane: The inner egg, or vitelline membrane, is the innermost membrane covering the egg mass. It is permeable to water and various dissolved substances, typically ions. During embryogenesis, the vitelline membrane incorporates proteins, non-protein nitrogen, carbohydrate, and salts in the chitinous part of the membrane (Pandian 1970). the outer egg membrane (chorion): The outer egg membrane (chorion) surrounds and protects the vitelline membrane, including the egg mass. Throughout various crustacean specimens (investigated so far), the chorion consists of three layers formed of epicuticular substances analogous to the crustacean cuticle (Morris and Afzelius 1967, Pandian 1970, Gilchrist 1978, Saigusa and Terajima 2000). It is made mostly of non-protein substances. To build and sustain an osmotic pressure within the egg, the chorion changes its permeability during embryogenesis and thus regulates the transmembrane flux of water and dissolved substances (Pandian 1970). crustacean embryo: a crustacean embryo is a developmental stage enclosed by the vitelline membrane and chorion of an egg. Embryogenesis constitutes the development of an individual in the part of the life cycle between fertilization and hatching. crustacean larva: A crustacean larva is in a postembryonic, free-​living, most often swimming developmental stage with its characteristic morphology. Larval development starts, by definition, with hatching or release from the vitelline membrane and proceeds often in a sequence of stages with the larva differentiating gradually, ending with a final molt into a juvenile. crustacean juvenile: The crustacean juvenile is in a developmental stage that has the final number of trunk segments and appendages resembling small adults, but is still not sexually mature. indirect development: Indirect development is a developmental mode in which a consecutive row of free-​living developing individuals (larvae) occurs between embryogenesis and the juvenile part of the life cycle. anamorphic development: Anamorphic development is a specific kind of indirect development. It typically starts with the hatching of an early larval form (nauplius or metanauplius larva) that gradually differentiates in a series of consecutive developmental stages. Growth takes place in a series of molts, with the larva adding and differentiating new segments and appendages until the juvenile phase is reached. Anostracans are an example taxon with anamorphic development. metamorphic (=hemianamorphic) development: Metamorphic development is another kind of indirect development. It implies the presence of an abrupt and dramatic body

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THE HATCHING MECHANISM The hatching mechanism in crustaceans depends on a number of different physiological and mechanical conditions. Hatching in crustaceans is not only triggered by the maturation of the developing individual at the end of embryogenesis. Various environmental factors such as diel, tidal, or lunar cycles; varying temperature; freezing; as well as various oxygen or osmotic conditions have also been shown to play crucial roles in initiating hatching (e.g., Pandian 1970, Forward 1987, Brendonck 1996, Charmantier and Charmantier-​Daures 2001). The hatching process can be divided into three main phases. The first phase includes osmotic rehydration and swelling of the embryo. The developing embryo and, in most crustaceans, the inner egg (vitelline) membrane start to swell by taking up water (resulting from an increasing osmotic pressure). The second phases consists of breakage of the outer egg membrane (chorion) caused by the swelling and increased pressure by the inner egg. The inner egg membrane is still intact at this stage and continues to envelop the hatchling. This has been reported in the majority of publications on crustacean development. The third phase includes the release of free, mobile larvae or juveniles. Termination of the hatching process is characterized by the dissolving or mechanical rupturing of the inner egg membrane and the final release of free larvae or juveniles. The inner egg membrane is thus also referred to as the hatching membrane in crustaceans. Hatching resulting from an increasing osmotic pressure within the developing egg, termed osmotic hatching (Needham 1931), is the most common hatching process among crustaceans; however, numerous exceptions exist. Figure 5.2 schematically depicts the hatching process of a hatching copepod nauplius larva passing through all three phases until hatching. During embryonic development, both egg envelopes (vitelline membrane and chorion) are highly impermeable, preventing the embryo from taking up water or losing solutes (e.g., Davis 1968, Morritt and Spicer 1995, Charmantier and Charmantier-​Daures 2001, Susanto and Charmantier 2001). At the end of embryogenesis, however, permeability of the egg envelopes changes and water is absorbed as a result of high osmotic pressure of the inner egg, which leads to swelling of the embryo and, finally, the burst or rupture of the chorion (e.g., Marshall and Orr 1954, Davis 1959, Pandian 1970, Saigusa and Terajima 2000, Charmantier and Charmantier-​Daures 2001). In decapod crustaceans, the water content can increase from 50% to 60% in freshly laid eggs to 70% to 80% in well-​developed prehatching eggs (Pandian 1970). The increase in permeability of the chorion at the end of embryogenesis coincides with the degradation of the chorion structure. This is caused by a production of proteolytic hatching enzymes during the prehatching period (e.g., De Vries and Forward 1991a,b, Geier and Zwilling 1998, Saigusa and Terajima 2000, Vogt 2008). It is known that in some brachyuran species, embryos release proteolytic enzymes at the onset of the hatching process (De Vries and Forward 1991b). These enzymes seemingly pass the vitelline membrane and degrade specific inner areas of the chorion, which finally results in its bursting [De Vries and Forward (1991b) and described later]. In addition, Saigusa and Terajima (2000) show that in the terrestrial crab Sesarma haematocheir, before rupturing of the chorion, the innermost chorion layer is digested

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Fig. 5.2. Illustrated hatching processes in Copepoda. Schematic illustration of the hatching process in the copepod Pseudocalanus minutus. From left to right:  embryo encapsulated within chorion and vitelline membrane, shortly before hatching (a); first bulging movements of the hatchling (b); bulging out of the chorion and emerging of the hatchling, still within the vitelline membrane (c, d); final release out of the vitelline membrane and free swimming nauplius larva (e). Modified from Marshall and Orr (1954).

by a caseinolytic protease, causing an inflow of water. In addition, in the crayfish Astacus astacus, an astacin-​like hatching enzyme has been identified. The expression level of this enzyme increases before hatching, whereas in newly hatched juveniles no such astacin-​like peptide or even transcripts are found (Geier and Zwilling 1998). Thus, the uptake of environmental water by the developing egg or embryo depends on regulation of the permeability of the egg membranes, along with an increasing osmotic egg pressure, which is caused by the embryos themselves (Pandian 1970, Charmantier and Charmantier-​Daures 2001). The osmolarity (and thus the osmotic pressure) inside the egg is regulated by the development of temporary or definitive embryonic osmoregulatory organs. Examples of the former include the nuchal glands or dorsal organ (e.g., the embryonic dorsal organ in the thermosbaenacean Tulumella unidens; Fig. 5.3A), whereas examples of the latter contain coxal or epipodite gills. Temporary osmoregulatory organs can be functionally replaced by definitive organs, usually gills, located on other parts of the body during late embryogenesis (Charmantier and Charmantier-​Daures 2001). Osmoregulatory organs in crustaceans are composed of ion-​transporting cells and also produce enzymes such as Na+/​K+-​adenosine triphosphatase or carbonic anhydrase, allowing the embryo to regulate the egg fluid osmolarity and to compensate for differences in salinity of the exterior habitat medium (e.g., Marshall and Orr 1954, Aladin and Potts 1995, Morritt and Spicer 1996, Charmantier and Charmantier-​Daures 2001). The capability of embryos to regulate the egg fluid osmolarity can also vary with the developmental strategies of crustaceans. It also differs with varying exposure to the external medium. Crustacean developing eggs can either be deposited “internally,” when the eggs are kept within a brood sac that may function as an osmotic protective incubation pouch (e.g., in the marsupium of isopods, in the brood sac of calanoid copepods, under the carapace of cladoceromorphans and thermosbaenaceans); or deposited “externally,” when the eggs are directly exposed to the surrounding medium (e.g., in decapods with eggs attached to the pleopods). The eggs may also be enveloped in a protective tertiary shell (resting egg or ephippium) and deposited in the soil or on hard substrata [e.g., the resting eggs of branchiopods and the egg strings of branchiurans (Fryer 1972, Charmantier and Charmantier-​Daures 2001, Hakalahti et al. 2004)]. With the ability of active osmoregulation, the embryo can absorb water to cause chorion rupture; however, the final hatching “event” of free-​swimming larvae or juveniles is the release from the vitelline membrane. Release occurs upon breakage of the vitelline membrane by enzymatic

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Fig. 5.3. Hatching processes in crustaceans. (A) Hatched larval stage of Tulumella unidens (Thermoscaenacea) and highlighted embryonic dorsal organ. Modified from Olesen et al. (2015). (B) Hatching phase of a Procambarus clarkii (Decapoda) juvenile by active movements of the appendages and pleon. Modified from Vogt (2008). (C) Hatched juvenile of P. clarkii, with chelipeds clamping the eggshell (arrowheads). Modified from Vogt (2008). (D) Backward release out of the chorion of a Petrolisthes armatus (Anomura) prozoea inside the vitelline membrane. Modified from Davis (1966c). (E–F) Backward hatching of a Euphausia pacifica nauplius (Euphausiacea) pushing out of the egg with its naupliar appendages. Modified from Gomez-Gutierrez (2002). (G) A nauplioid larval stage of Americamysis bahia (Peracarida, Mysidacea) that hatches by straightening the pleon and rupturing the vitelline membrane. Modified from Wortham-Neal and Price (2002). (H) Hatched nauplioid stage of Praunus inermis surrounded by a nauplioid cuticle. Modified from San Vicente et al. (2014). (I) Prehatching stage of Anaspides tasmaniae (Anaspidacea) in the vitelline membrane. Burst chorion halves are still attached to the sphere of the vitelline membrane. Modified from Hickman (1936).

dissolution or by mechanical forces such as pleonal movements of the mother animal, appendage or abdominal movements of the hatchlings (e.g., puncturing the membrane with cuticular spines on the dorsal urosome in peracarids), or by biting with an embryonic “egg tooth” as in amphipods (Pandian 1970, Forward and Lohmann 1983, Forward 1987). In addition, during breakage and larval release in decapod crustaceans, a cocktail of specific dipeptides (a mixture of arginine and glycine) appears to be present in the water column. These dipeptides are detected by peptide-​specific receptors of the mother animal and this invokes a stereotypic parental pleonal “pumping” behavior (Rittschof et al. 1985, Forward 1987). Chemical stimulation of the hatching process is also present

Hatching in the barnacle Semibalanus balanoides (Cirripedia). In this species, a hatching substance or egg-​ hatching pheromone [potentially a product of the eicosapentaenoic acid biosynthesis (see Clare 1997)] is secreted by the mother animal into the mantle cavity, which activates hatching of nauplius larvae [in contrast to what is found in decapods; see “Decapoda” under “Hatching Features” this chapter (Crisp 1956, 1969, Crisp et al. 1991, Anderson 1994)]. The final release of larvae or juveniles occurs either simultaneously with chorion rupture or shortly after, usually after the animals have spent time outside the chorion but still within the vitelline membrane. After ripping the chorion halves, some copepod and branchiopod hatchlings are free floating and can further differentiate within the vitelline membrane before they are finally released (e.g., Davis 1959, Belk 1987, Fryer 1996, Fritsch and Richter 2015).

HATCHING FEATURES IN CRUSTACEANS Decapoda and Stomatopoda Within the decapods, there is a large variation in body forms. Nevertheless, most decapods show a metamorphic developmental pattern, and the developing eggs are carried and protected by the mother animal (e.g., in Caridea, Stenopodidea, Reptantia, Amphionidacea). In this group, the majority of species spawn their eggs into the water column and hatchlings are released as free-​ swimming larvae (Scholtz 2000, Vogt 2013). In decapods with brood care, eggs are attached and carried by the pleopods [e.g., caridean shrimps, lobsters, and crabs; rarely, eggs are carried by the last pairs of pereopods, such as in Lucifer faxoni (Lee et al. 1992)]. A notable exception is the group of Dendrobranchiata, most of which spawn their eggs in the water column and hatchlings develop through a consecutive row of free-​swimming nauplius, protozoea, and mysis larvae. Except for the previously mentioned dendrobranchiates, most decapods and stomatopods proceed through an embryonic “egg–​nauplius” stage and hatch as a more developed larva (e.g., of the zoea-​type, protozoea, mysis, phyllosoma) with additional segments and functional thoracopods, or as decapodid larva (e.g., megalopa) with functional thoracopods and pleopods (Scholtz 2000, Anger 2001, Vogt 2013, Jirikowski et al. 2013, 2015). At the end of embryonic development, the yolk mass within the egg may be completely consumed, and the prehatchlings show distinctive stretching and levering appendage movements (Fig. 5.3B–​D; Forward 1987, Vogt 2008). Within the embryonic cephalic region, a well-​developed pair of compound eyes may be present [except in developing dendrobranchiates (Davis 1966c, Pandian 1970, Forward 1987, Vogt 2008)]. It has been suggested that the visual system becomes functional just before the event of hatching to perceive environmental clues (Forward 1987). Hatching or larval release in most decapods does not occur randomly; it is quite precisely scheduled and depends upon exposure to several environmental factors such as diel, tidal, or lunar cycles (Ennis 1973, Forward and Lohmann 1983, Forward 1987). In particular, in brachyurans, the diel, tidal, and lunar cycles play a crucial role in the timing of hatching (see Chapter 17 in volume 6). Often, the release of larvae occurs at night and is synchronized with the time of high tide (Saigusa 1992a,b, 1993, 2002, Anger et al. 2015). Experiments in the crab Sesarma haematocheir revealed that the timing of hatching seems to be endogenously controlled and it is induced by the mother animal 48 to 49.5 hours before larval release [see also Figure 12 in Saigusa (1993)]. Once this stimulus, the so called hatching-​program inducing factor (HPIF), is released by the mother animal, an endogenous clock of the embryos determines the hatching process and time (Saigusa 1992a,b, 1993, Ikeda et al. 2006). HPIF acts as an important key stimulus for the circatidal rhythm in larval release. The critical time frame of inducing hatching often corresponds to the time of high tide two nights before hatching (Saigusa 1992a).

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Developmental Biology and Larval Ecology Environmental factors not only induce the hatching process, but also may have positive effects on the hatching or larval survival rate. For instance, crab species living near coastlines can release and distribute their larvae quickly and more widely during high tides. Hatching of species that live preferably in the intertidal zone occurs near the maximum-​amplitude high tides, so that the offspring can be transported offshore quickly (Anger et  al. 1994, Morgan and Christy 1995). Thus, visual predators can be avoided most effectively if larvae are released on nocturnal maximum high tides (Christy and Stancyk 1982, Salmon et  al. 1986, Forward 1987). The timing of larval release ultimately can depend on whether adults, embryos, or larvae are most vulnerable to predators. If females or embryos are more at risk, then crabs will remain near their refuges to release larvae. The safest time for intertidal crabs to release larvae is at the beginning of the night when many planktivorous fish do not feed (Morgan and Christy 1995, Anger et al. 1994, Anger et al. 2015). The hatching time of coastal brachyurans is partly driven by tidal schedules, and the larval release seems indirectly coordinated with low abundance of planktivorous predators (Christy 2011). In most decapods, the release of larvae can last for only a few minutes and may be assisted by rapid pleon movements of the mother animal. It can also last for several days or even a month (e.g., Branford 1978, Forward and Lohmann 1983, Kunisch and Anger 1984, Forward 1987, De Vries and Forward 1991a). In littoral crabs of the genus Sesarma, Uca, or Petrolisthes, or in the lobster Homarus gammarus, a particular hatching behavior is observed. Female crabs elevate their body and pump their pleon vigorously back and forth to produce strong water currents. During hatching and breakage of eggshells, a heterogeneous mixture of dipeptides (arginine and glycine) is released into the water (it is not specifically known whether this mixture is directly produced by the embryos). These dipeptides are detected by the mother animal via specific peptide receptors and provoke or enhance the female “pumping” behavior. Pleonal pumping causes more eggs to hatch by mechanically rupturing the vitelline membrane, which in turn increases the dipeptide concentration and causes further pumping movements until all larvae or juveniles have hatched (e.g., Branford 1978, Forward and Lohmann 1983, Rittschof et al. 1985, Forward 1987). Although parental assistance during hatching appears to be common in decapods—​for example, females of the anomuran crab Petrolisthes violaceus use their fifth pereopods to aid in larval release (Förster and Baeza 2001)—​ mechanical hatching support is not essential for larval release. Eggs from various decapod species such as the subtidal crab Rhithropanopeus harrisii or the lobster Homarus gammarus also hatch normally, even when the eggs are removed from the mother animal (Branford 1978, Forward and Lohman 1983, Vogt 2008). Hatching larvae open up the vitelline membrane mechanically with their appendages (thoracopods and or pleopods) or by straightening their pleon (e.g., Davis 1966a, Forward 1987, De Vries and Forward 1991a, Vogt 2008). In sum, although larval egg membrane release may be supported mechanically by the mother animal, the specific hatching time seems to be controlled or induced by the embryo (Forward and Lohman 1983, De Vries and Forward 1991b, Vogt 2008). Euphausiacea Euphausiid crustaceans show a metamorphic developmental mode and mostly release their eggs into the pelagial. They use a free-​floating egg strategy, but developing eggs can also be carried and protected by the mother animal (e.g., Nematoscelis difficilis) when attached to the posterior thoracopods in a membranous sac (Mauchline and Fisher 1969, Gomez-​Gutierrez 2003). The offspring of free-​floating eggs hatch as orthonauplius larvae, whereas metanauplius (or pseudo-​ metanauplius) hatch from the membranous sac (Fig. 5.3E, F; Gomez-​Gutierrez 2002, 2003, Ambriz-​ Arreola et al. 2015, Akther et al. 2015). Detailed hatching information on Euphausiacea is limited, but the available hatching data of the euphausiids Euphausia pacifica, Thysanoessa spinifera, T.  inspinata, and Nematoscelis difficilis

Hatching show some small differences from other crustaceans. Shortly before hatching, the emerging embryo turns yellowish and there is no particular embryo growth or size increase caused by increasing osmotic pressure. Furthermore, before chorion breakage, the vitelline membrane is ruptured by movements of the embryo [also defined as the twitching stage (Ambriz-​Arreola et al. 2015)]. When ruptured, the rest of the vitelline membrane and the nauplius remain freely suspended within the chorion. To hatch, hatchlings use their naupliar appendages to press their abdomen backward, break up the shell, and squeeze out of the chorion—​termed backward hatching (Fig. 5.3F). The presence of enzymatic dissolution or weakening of the chorion structure during the hatching process has not been convincingly verified (Gomez-​Gutierrez 2002). Euphausiids with an extended embryonic phase hatch as a metanauplius or a pseudo-​metanauplius, or in the calyptopis I stage. In contrast to the orthonauplius stage, these hatchlings release themselves from the chorion, often in a forward direction, by flipping or pushing off the chorion halves using their abdomen or appendages (Gomez-​Gutierrez 2002, 2003). Peracarida Peracarid crustaceans develop directly and show an extremely diverse adult morphology—​from the laterally flattened amphipods to the dorsoventrally depressed isopods and tanaidaceans; the thermosbaenaceans, spelaeogriphaceans, and mictaceans; to shrimp-​like lophogastrids and mysids; and the comma-​shaped cumaceans. These taxa share the possession of a female brood pouch (marsupium) formed by the oostegites (spoon-​shaped plates) of the thoracopods [except Thermosbaenacea, which have a dorsal brood pouch (Olesen et al. 2015)]. Eggs are released into the incubating brood pouch medium, which—​at least in some taxa—​is thought to provide additional nutrition to the developing embryos. The fluid or medium probably also serves as an osmotically stable environment to optimize growth conditions (e.g., Morritt and Spicer 1996, Morritt and Richardson 1998, Charmantier and Charmantier-​Daures 2001). Free-​swimming pelagic larvae in peracarids are only known for parasitic gnathiid isopods such as Gnathia maxillaris (Hispano et al. 2014). Most other peracarid larvae hatch or are released from the egg membranes within the brood pouch as a “manca” larva that closely resembles small juveniles or adults but lacks the last thoracopods compared to the adult form (Martin 2014). Amphipod and isopod eggs are laid directly into a semiclosed ventral marsupium where they are fertilized and pass through the embryonic stages. The number of egg membranes in amphipods and isopods is still not completely clear. At least one egg membrane, which is considered the vitelline membrane, has been described in several species (e.g., Davis 1964, Sheader and Chia 1970, Morritt and Spicer 1995, 1996). The terrestrial woodlouse Porcellio scaber carries its developing embryos in the protected fluid-​filled marsupium. Eggs of P. scaber clearly have an outer chorion and an inner vitelline membrane (Wolff 2009). Before hatching, the osmotic inflow of water occurs, which is controlled via the permeability of the vitelline membrane and by activity of the dorsal organ in the developing embryo. Late-​embryonic-​stage organisms are temporarily able to regulate the osmolarity of the periembryonic fluid [the space between the embryo and the vitelline membrane (e.g., Morritt and Spicer 1995, 1996, Charmantier and Charmantier-​Daures 2001]. At the end of embryonic development, the vitelline membrane is ruptured by trunk or appendage movements and by puncturing the membrane, facilitated by cuticular spines on the dorsal urosome of the hatchling (Davis 1964, Sheader and Chia 1970). After hatching, the individuals are still enclosed in the protective marsupium and develop, commonly involving one or more molts, until they are finally released by the mother animal. As all other peracarids, mysids lay their eggs into the ventral marsupium, and embryonic development entirely takes place within the protective brood pouch. Release of the free-​swimming larvae from the marsupium usually occurs at night (Wortham-​Neal and Price 2002). Within the brood

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Developmental Biology and Larval Ecology pouch, three consecutive developmental phases occur. The first phase (also called the embryonic phase) ends with rupturing or shedding of the chorion resulting from osmotic water uptake and straightening of the rudimental pleon of the embryo (Fig. 5.3G; Mauchline 1980, Wortham-​Neal and Price 2002). The embryo at this stage is in the process of germ band elongation, and musculature forms only later during the postnauplioid stage ( Jirikowski et al. 2013). Hence, the straightening of the pleon is performed by a nonmuscle-​driven shape change of the embryo. In general, this is considered the hatching event. The following developmental stages (Fig. 5.3H, nauplioid and postnauplioid stages) that are still yolk dependent remain immobile and are enclosed by a membrane called the nauplioid cuticle that, in contrast to the adult cuticle, does not lie directly on top of the epidermis of the embryo. It is unclear whether this membrane represents a vitelline membrane or an embryonic cuticle. After this envelope is shed, the postnauplioid-​stage embryos differentiate until the yolk mass is resorbed. Eventually, swimming juveniles are released from the female marsupium (Wortham-​Neal and Price 2002). Branchiopoda The mode of reproduction in branchiopod crustaceans is extremely complex and depends on their life cycle. Two types of eggs can be produced: subitaneous eggs in which embryos continue development until hatching as pseudo-​direct larvae or juveniles (predominantly in Cladoceromorpha, but also present in some anostracan groups), or resting eggs with a tertiary shell (“third” egg shell) that host a diapaused gastrula stage in which the animal is capable of hibernation through periods of harsh environmental conditions (e.g., Flößner 1972, Fryer 1996, Dumont and Negrea 2002, Fritsch et al. 2013). Subitaneous eggs are mostly generated during parthenogenetic reproduction and mature within the ovaries (e.g., Martin 1992, Olesen 1999, Dumont and Negrea 2002, Fritsch et al. 2013). Subsequently, these eggs can be deposited either into brood pouches that are attached to the genital segments (e.g., in Anostraca as Artemia parthenogenetica) or placed and carried directly under the dorsal carapace, which is often modified to form a brood chamber [in cladoceromorphans (Egloff et  al. 1997, Kotov and Boikova 1998, Rivier 1998, Olesen 1999, Flößner 2000)]. Hatching from subitaneous eggs occurs when free-​swimming nauplius larvae are released from the anostracan brood pouch or pseudo-​direct developing larvae are carried in cladoceromorphans in the brood chamber under the carapace. The brood chamber may have a nourishing role (Nährboden) during development [e.g., in Onychopoda, Penilia, and Moina spp. (Egloff et al. 1997, Dumont and Negrea 2002)]. Subitaneous developing embryos are enclosed by two egg membranes or shells (Kotov and Boikova 1998, 2001, Fritsch and Richter 2012). In contrast to resting eggs, these membranes are more elastic and less chitinous (Kotov and Boikova 1998, 2001). Branchiopod resting eggs are mostly produced by gamogenesis and are discarded in the soil of temporary dried-​out water ponds or pools (Brendonck and De Meester 2003, Alekseev et al. 2007, Fritsch et al. 2013). Hatching of resting eggs is induced or activated by distinct abiotic factors such as photoperiod, illumination, temperature, oxygenation, mineralization, osmolarity, and desiccation. In some branchiopods hatching of resting eggs is influenced by a number of key factors, such as freezing period, refilling of temporary pool, increase in the water level, and the size or quality of the egg bank [e.g., Eubranchipus gelidus, Triops longicaudatus, Daphnia (see Pancella and Stross 1963, Scott and Grigarick 1979, Vanhaecke and Sorgeloos 1989, Vanderkhove et al. 2005, Vanschoenwinkel et al. 2010)]. Hatchlings from resting eggs are released either as free-​swimming nauplius larvae in anostracans, notostracans, laevicaudatans, and spinicaudatans (Fig. 5.4A, all examples of anamorphic developmental patterns); or as juveniles in the cladoceromorphans that have direct development [Fig. 5.4B, except from resting eggs of Leptodora kindtii (Olesen et al. 2003, Olesen 2004)]. The ephippium (dorsal carapace section that encloses resting eggs) of cyclestherids and anomopod

Hatching (A)

(D)

(B)

(E)

(C)

(F)

Fig. 5.4. (A) Hatching nauplius of Artemia franciscana (Branchiopoda, Anostraca) (original by Thomas Frase). (B) Prehatching stage of Cyclestheria hislopi (Branchiopoda, Cyclestherida). Burst chorion halves are still attached to the sphere of the vitelline membrane. (C) Clutch of developing embryonic stages of Argulus japonicus (Branchiura). (D) Release of an embryo of A. megalops and its surrounding vitelline membrane from the chorion. Modified from Davis 1966b. (E, F) Embryos of Tigriopus californicus (Copepoda, Harpacticoida) at the beginning of hatching. The embryos are still contained in the vitelline membrane, but have been released from the egg sack compartments. A1, antennula; A2, antenna; ca, carapace; ch, chorion; el, eye lobe; Ex, exopod; MD, mandible; vm, vitelline membrane.

cladocerans also functions as a protective and preservative tertiary shell and is thus normally also considered a sort of resting egg (Roessler 1995, Hiruta and Tochinai 2014, Fritsch and Richter 2015). The tertiary egg shell or the cyst case of branchiopod resting eggs not only protects the diapausing embryo from mechanical and radiation damage (Belk 1972, Fryer 1972, 1996, Korovchinsky and Boikova 1996), but also it preserves membrane integrity during the rehydration process. In addition, an accumulation of glycerol within the tertiary shell facilitates osmotic water absorption after the embryo has been reactivated (Brendonck 1996, Dumont and Negrea 2002). The dormant gastrula or early embryo within the resting egg is likely reactivated by the transport of calcium ions through the egg membranes (Dumont et  al. 1992). This activates calmodulin and dormant kinase enzymes, which restart the “cellular metabolism” of the gastrula. To build up or increase the osmotic pressure in the resting egg, osmotically active substances are accumulated within the embryonic fluid (Belk 1972). After further development and swelling of the embryo and vitelline membrane, the tertiary eggshell and chorion finally crack and the embryo, still within the vitelline membrane, is released (e.g., Morris and Afzelius 1967, Mossin 1986, Belk 1987, Onbé 1991, Mugrabe et al. 2007). The vitelline membrane spheres of cyclestherids and anomopods tend to be attached

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Developmental Biology and Larval Ecology to the opened ephippium (Egloff et al. 1997, Fritsch and Richter 2015). Before the final release of the hatchling, the vitelline membrane can expand tremendously and, within this “container,” the free-​floating, developing prehatching nauplii or juveniles are quite active (Fryer 1972, Roessler 1995, Fryer 1996, Fritsch and Richter 2015). Branchiura Branchiurans normally show a metamorphic developmental pattern (Olesen 2018). These parasitic crustaceans spend most of their life attached to their fish hosts and live primarily in fresh water. Their eggs are deposited away from the host and are commonly laid in strings or batches in one or more rows, in single or multiple layers on smooth plant or hard substratum surfaces (Fig. 5.4C; e.g., Davis 1966b, Shafir and van As 1986, Overstreet et al. 1992, Hakalahti et al. 2004, Møller 2009). The eggs are coated with a gelatinous compound that hardens after the eggs are laid (Davis 1966b, Overstreet et al. 1992, Møller 2009, Møller and Olesen 2014). Embryonic development and hatching of branchiurans are strongly correlated to the water temperature and a specific photoperiod, and range from at least 12 to 81 days (e.g., Shafir and van As 1986, Overstreet et al. 1992, Rushton-​Mellor and Boxshall 1994, Møller and Olesen 2014). Developing embryos are enclosed in a very delicate vitelline membrane and a thick, yellow-​brownish chorion. Hatching in branchiurans is caused by osmotic swelling (Davis 1966b). At the end of embryonic development, the chorion ruptures and hatching larvae release themselves actively (Fig. 5.4 D). After release of the larvae, the vitelline membrane remains partially within the chorion (Davis 1966b). Three different types of hatching larvae are known for branchiurans. A metanauplius-​like (e.g., Argulus foliaceus), a juvenile-​ like (e.g., A. megalops), and a non-​swimming type of larvae [e.g., Chonopeltis brevis (Fryer 1961)]. However, most of the current knowledge on branchiuran hatching is based on observations of Argulus species (Møller and Olesen 2014). Thecostraca Members of the Thecostraca show a well-​developed metamorphic development pattern and the group comprises three major lineages: Facetotecta, Ascothoracida, and Cirripedia. Larval development in all three taxa has been studied extensively; however, information on the hatching process is limited. Thecostracans hatch either as a nauplius or as a well-​developed cyprid (a-​, c-​, or y-​cyprid) larva (Høeg et al. 2014). Lepadomorph and balanomorph barnacles (Cirripedia) release fertilized eggs into their mantle cavity and the eggs are combined in an egg mass. The eggs are then either attached to ovigerous frena (tegumentary folds) or directly on the surface of the inner mantle cavity (Walker 1983). Embryonic development takes place entirely within the mantle cavity. Data about osmotic swelling of the embryo or the vitelline membrane and final rupture of the chorion are not available for cirripeds. Before hatching of the nauplius or cyprid larvae, a “hatching substance” is released by the adults to promote larvae to hatch (Crisp 1956, 1969). This hatching substance is a derivative of eicosapentaenoic acid biosynthesis (Clare et al. 1985) and it is produced within the hypodermis mantle cavity layer of the mother animal (Holland 1987). Copepoda Copepod crustaceans generally show a metamorphic development pattern and an extraordinarily high variability of life strategies and body shapes, including free-​living small planktonic forms and vermiform-​like species that are widely distributed in interstitial habitats (Olesen 2018). They can be associated with different substrata and are often parasites on almost all aquatic metazoans. The

Hatching copepod life cycle generally consists of an embryonic phase followed by a naupliar and a copepodid phase with a before characteristic larval morphology (Boxshall 1992, Huys 2014). Besides having various modes of life and disparate morphology, copepods have also evolved an extremely diverse range of egg-​carrying methods. Their eggs can be carried in a protective and nourishing paired or unpaired egg sac. Sometimes, they are surrounded by a secreted mucus egg mass attached to the ovigerous spines. In other (pelagic) species, the eggs are packed in linear “strings” attached to the genital appendages or segments, or are connected by axial filaments to the genital aperture. The eggs can also be laid directly into the host animal or simply liberated into the water column (Huys 2014). Before hatching of copepod nauplius larvae, osmotic pressure within the developing egg starts to increase and the embryo swells by absorption of water. In calanoid copepod eggs, the increasing fluid osmolarity is regulated by salt secretion or excretion of the embryo (Marshall and Orr 1954, Davis 1959). The volume increase in eggs freely released into the water is much greater than in eggs that are carried. The chorion first bursts and the embryo within the vitelline membrane bulges out. Copepod embryos mechanically aid chorion rupture by twitching and stretching their body. In addition, they also use their antennae to puncture the vitelline membrane (Marshall and Orr 1954, Davis 1959). Emerging embryos enclosed within the vitelline membrane are completely detached from the chorion and become more and more active within the expandable vitelline membrane [e.g., Calanus finmarchicus (Marshall and Orr 1954)]. The vitelline membrane finally ruptures or is actively torn away by the hatching nauplius larva. Hatchlings within an egg sac are released in a similar manner, but the vitelline membrane is never completely detached from the chorion. In species such as the harpacticoid Tigriopus californicus, the egg sac is organized like a beehive, with each egg in a separate compartment. This structure dissolves when hatching is initiated (Fig. 5.4E, F). In addition, hatching from an egg sac can be supported by the movements by the mother animal. These hatchlings may hatch almost synchronously (Marshall and Orr 1954, Davis 1959).

DEVELOPMENTAL MODE AND TIMING OF HATCHING The events that constitute hatching vary with the developmental patterns [anamorphic, metamorphic (=hemianamorphic) or direct (=epimorphic) development], the extent of maternal brood care, and the life history strategy of crustaceans. Figure 5.5 shows how specific developmental processes are timed relative to hatching. The developmental processes considered here are embryonic swelling, vitelline membrane and chorion rupture, and body movement of the hatchling. The relative timing is displayed for a selection of representatives with various developmental modes that have been introduced in the previous sections: anamorphic development: Artemia sp. (Anostraca); metamorphic development: Euphausia pacifica (Euphausiacea), Palaemonetes varians (Decapoda); and epimorphic (direct) development:  Daphnia magna (Branchiopoda), Americamysis bahia (Mysidacea), and Porcellio scaber (Isopoda). Embryonic swelling generally precedes shedding of the chorion and vitelline membrane by a short period (Fig. 5.5). The relative timing of the remaining developmental processes is less conserved. The vast majority of crustaceans with an anamorphic development pattern (e.g., Artemia sp., Fig. 5.5) hatch as free-​swimming larvae and become fully active once liberated from the egg envelopes. Thus, the hatching processes (embryonic swelling, removal of the chorion, and removal of the vitelline membrane) are naturally preceded by morphogenesis and organ differentiation of the nauplius larva. Morphogenesis and organogenesis continue throughout the larval phase of the life cycle (Fig. 5.5). In the branchiopod example, the chorion and vitelline membrane are mostly shed simultaneously. However, within anamorphically and metamorphically developing

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Fig. 5.5. Schematic overview of embryonic development and hatching processes in relation to crustacean developmental modes as represented by six species. All developmental phases (i.e., morphogenesis, organ differentiation, body movement, free-​living individual) and the hatching-​related events (i.e., embryonic swelling, chorion rupture, vitelline membrane rupture) are illustrated in schematic timelines. The six species were selected to cover a significant diversity in crustacean developmental modes:  Artemia sp. (anamorphic development); Euphausia pacifica and Palaemonetes varians (metamorphic development); and Daphnia magna, Americamysis bahia, and Porcellio scaber (direct development). The morphology of the hatchling (nauplius, zoea, juvenile), the developmental mode (indirect anamorphic development, indirect metamorphic development, and direct development), as well as the type of brood care (external, semi-​internal or absent) are indicated in the columns on the left.

Hatching crustaceans, there are variations in the timing of chorion and vitelline membrane shedding, as demonstrated when comparing branchiopod and euphausiacean hatching (Fig. 5.5). In the latter example, the vitelline membrane is shed before the chorion. Thus, for a certain period during development, the nauplius larva is active while still being enclosed in the large chorion. Representatives with metamorphic development but external brood care, such as caridean shrimp (e.g., Palaemonetes varians; Fig. 5.5), share a similar timing pattern of hatching-​related processes with anamorphically and metamorphically developing species. Decapods, as well as copepods, various branchiopods, and euphausiids, show extensive naupliar appendage or trunk body movement caused by controlled muscular activity in the prehatching stage prior to the shedding of the chorion and vitelline membrane. Taxa with epimorphic development (=direct development) and semi-​internal brood care (in chambers/​pouches formed by limbs or carapace, exemplified by cladoceromorphans with subitaneous eggs, mysids, and isopods in Fig. 5.5), however, show a clear tendency to separate chorion and vitelline membrane shedding. Some copepods and branchiopods also shed their chorion first, and the prehatching stages remain within the expanded vitelline membrane until final hatching. Rupture of the chorion in these representatives occurs significantly earlier than rupture of the vitelline membrane (i.e., before morphogenesis and organogenesis in the embryo are completed). The most extreme case is observed, however, in mysids, which hatch from the chorion while the embryo is still in the process of developing the body segment primordia and before musculature or nervous system development (see Jirikowski et al. 2013). In mysids, the majority of embryonic development is carried out after the first “hatch.” Furthermore, in species with semi-​internal brood care, final hatching from the vitelline membrane precedes release of juveniles from the brood chamber. It is thus the liberation from the marsupium and not the hatching event that marks the beginning of the free-​living part of the life cycle in these species. However, some mysid species molt to the fully mobile juvenile stage only shortly after they have been released from the marsupium (Wortham-​Neal and Price 2002). In the case of some branchiopods (e.g., Cyclestheria), juveniles molt several times before leaving the chamber (Roessler 1995, Olesen 1999, Fritsch and Richter 2012). Apparently, the life history strategy of semi-​internal brood care has a strong influence on the timing of hatching-​related developmental processes. It is likely that the brood pouch physiologically “replaced” the chorion during evolution of these lineages, providing an appropriate milieu for both embryonic development and mechanical protection. This applies to a large number of crustaceans, such as Peracarida and Cladoceromorpha. All these groups lack a primary free-​living larval phase (except for cladoceran haplopod resting egg hatchlings). Taken together, the hatching procedure in crustaceans is linked strongly to the life history strategy in terms of timing.

A CLASSIFICATION OF HATCHING MODES In the previous sections we demonstrated the diversity of hatching processes across crustacean taxa. The hatching process itself comprises various events before to hatching:  enzymatic degradation of the envelopes, swelling of the embryo by water uptake, swelling of the vitelline membrane, embryonic shape change, active movement of the appendages or the trunk, and squeezing or pushing of the soft body through a ruptured hole in the enveloping membranes or puncturing of these with a specialized structure as seen in amphipods, euphausiaceans, or anostracans. Certain species may lack many of these prehatching processes, and none of them can cause hatching alone. Combinations of different prehatching processes are necessary for successful hatching, and their combination varies throughout the Crustacea. Here we summarize several combinations of prehatching processes, which we refer to as characteristic hatching modes.

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Developmental Biology and Larval Ecology Mode 1: Appendage-​mediated Hatching This mode applies to crustaceans that have either anamorphic or metamorphic development and hatch as a nauplius larva. The nauplius larva is fully differentiated at the end of embryogenesis and begins the hatching process with flexing and extension movements of the naupliar appendages, thereby adding to the mechanical stress on the enveloping membranes (enzymatic degradation of the chorion and embryonic swelling also facilitate rupture). During membrane release, the hatchling often squeezes or pushes itself through an opening in the envelopes. Appendage-​mediated hatching representatives include anostracans, notostracans, laevicaudatans, spinicaudatans, resting egg copepods, thecostracans, branchyurans, and euphausiaceans. Mode 2: Trunk-​mediated Hatching This mode is generally found in metamorphic crustaceans that hatch as an advanced larva with a well-​developed and movable posterior trunk (zoea-​like larvae), or in epimorphically developing crustaceans that hatch as juveniles. Enzymatic degradation and embryonic swelling co-​occurs, but the individual is liberated from the envelopes by extension movements of the pleon. Maternal assistance is also commonly found in these taxa, which involve particular movements of the mother animal to assist the breakage of the envelopes mechanically. The late embryos may also have specialized cuticular structures and high rigidity to penetrate the egg envelopes actively (e.g., in crayfish). Trunk-​mediated hatching representatives include amphionidaceans, anaspidaceans, carideans, brachyurans, anomurans, acheleatas, stenopodideans, and stomatopods. Mode 3: Extended Hatching Many species with semi-​internal brood care hatch from the chorion during embryogenesis and continue developing within the vitelline membrane or an embryonic cuticle in the mother’s brood pouch. Embryonic swelling, enzymatic degradation, and shape change of the embryo contribute to the rupture of the chorion, but active muscle-​generated movement does not play the same role as in modes 1 and 2. The vitelline membrane or embryonic cuticle is shed later during development, and under the influence of muscular contraction in the embryo. Extended hatching representatives include cladoceromorphans (except the resting egg of haplopod hatchlings), leptostracans, and peracarids.

SUMMARY We presented here the first attempt to provide a comparative overview of the hatching process in crustaceans and made efforts to address the diversity of this process. Hatching is a multifaceted process during which the physiological, ecological, and evolutionary aspects of development are reflected. A common denominator of hatching in all crustaceans is embryonic swelling by active water uptake. The remaining processes and subprocesses vary among taxa. Our classification of hatching modes is intended as a starting point for a more inclusive approach to the hatching phenomenon in a group as diverse and disparate as crustaceans. Thus, it should also be viewed as a preliminary attempt and, hopefully, subject to future completion and modification as knowledge of crustacean development continues to grow.

Hatching

ACKNOWLEDGMENTS Georg Brenneis, Gerhard Scholtz, and Carsten Wolff are thanked for fruitful discussions in the progress of preparing this chapter. We also thank Thomas Frase for providing hatching branchiopod image data.

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Hatching Olesen, J. 2014b. Cladocera: Anomopoda. Pages 63–​68 in J.W. Martin, J. Olesen, and J.T. Høeg, editors. Atlas of Crustacean Larvae. Johns Hopkins University Press, Baltimore, Maryland. Olesen, J. 2014c. Cyclestherida. Pages 58–​62 in J.W. Martin, J. Olesen, and J.T. Høeg, editors. Atlas of Crustacean Larvae. Johns Hopkins University Press, Baltimore, Maryland. Olesen, J. 2018. Crustacean life cycles: developmental strategies and environmental adaptations. Pages 1–​34 in G. Wellborn, and M. Thiel, editors. The Natural History of the Crustacea. Volume 5: Life Histories. Oxford University Press, New York, New York. Olesen, J., T. Boesgaard, and T.M. Iliffe. 2015. The unique dorsal brood pouch of Thermosbaenacea (Crustacea, Malacostraca) and description of an advanced developmental stage of Tulumella unidens from the Yucatan peninsula (Mexico), with a discussion of mouth part homologies to other Malacostraca. PLoS One 10:e0122463. Olesen, J., and S. Richter. 2013. Onychocaudata (Branchiopoda: Diplostraca), a new high-​level taxon in branchiopod systematics. Journal of Crustacean Biology 33:62–​65. Olesen, J., S. Richter, and G. Scholtz. 2003. On the ontogeny of Leptodora kindtii (Crustacea, Branchiopoda, Cladocera), with notes on the phylogeny of the Cladocera. Journal of Morphology 256:235–​59. Onbé, T. 1991. Some aspects of the biology of resting eggs of marine cladocerans. Pages 41–​55 in A. Wenner and A. Kuris, editors. Crustacean Egg Production. Crustacean Issues. Volume 7. A.A. Balkema, Rotterdam, The Netherlands. Overstreet, R.M., I. Dykova, and W.E. Hawkins. 1992. Branchiura. Pages 385–​413 in F.W. Harrison and A.G. Humes, editors. Microscopic Anatomy of Invertebrates. Volume 9. Crustacea. Wiley-​Liss, New York, New York. Pancella, J.R., and R.G. Stross. 1963. Light induced hatching of Daphnia resting eggs. Chesapeake Science 4:135–​140. Pandian, T.J. 1970. Ecophysiological studies on the developing eggs and embryos of the European lobster Homarus gammarus. Marine Biology 5:154–​167. Punzo, F. 2000. Postembryonic development. Pages 47–​74 in F. Punzo, Olesen, editors. Desert Arthropods: Life History Variations. Springer, Berlin, Germany. Rittschof, D., R.B. Forward Jr., and D.D. Mott. 1985. Larval release in the crab Rhithropanopeus harrisii (Gould): chemical cues from hatching eggs. Chemical Senses 10:567–​577. Rivier, I.K. 1998. The Predatory Cladocera (Onychopoda: Podonidae, Polyphemidae, Cercopagidae) and Leptodorida of the World. Backhuys, Leiden, The Netherlands. Roessler, E.W. 1995. Review of Colombian Conchostraca (Crustacea): ecological aspects and life cycles: family Cyclestheriidae. Hydrobiologica 298:113–​124. Roonwal, M.L. 1944. Some observations on the breeding biology, and on the swelling, weight, water-​content and embryonic movements in the developing eggs of the Moluccan king-​crab, Tachypleus gigas (Müller) [Arthropoda, Xiphosura]. Proceedings of the Indian Academy of Sciences B: Biological Sciences 20:115–​129. Rushton-​Mellor, S.K., and G.A. Boxshall. 1994. The developmental sequence of Argulus foliaceus (Crustacea: Branchiura). Journal of Natural History 28:763–​785. Saigusa, M. 1992a. Control of hatching in an estuarine terrestrial crab I: hatching of embryos detached from the female and emergence of mature larvae. The Biological Bulletin 183:401–​408. Saigusa, M. 1992b. Observations on egg hatching in the estuarine crab Sesarma haematocheir. Pacific Science 46:484–​494. Saigusa, M. 1993. Control of hatching in an estuarine terrestrial crab II: exchange of a cluster of embryos between two females control of hatching in an estuarine terrestrial crab II: exchange of a cluster of embryos. The Biological Bulletin 184:186–​202. Saigusa, M. 2002. Hatching controlled by the circatidal clock, and the role of the medulla terminalis in the optic peduncle of the eyestalk, in an estuarine crab Sesarma haematocheir. Journal of Experimental Biology 205:3487–​3504. Saigusa, M., and M. Terajima. 2000. Hatching of an estuarine crab, Sesarma haematocheir: from disappearance of the inner (E3) layer to rupture of the egg case. The Journal of Experimental Zoology 287:510–​523. Salmon, M., W. Seiple, and G. Morgan. 1986. Hatching rhythms of fiddler crabs and associated species at Beaufort, North Carolina. Journal of Crustacean Biology 6:24–​36.

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6 PATTERNS OF LARVAL DEVELOPMENT

Ole Sten Møller, Klaus Anger, and Guillermo Guerao

Abstract In this chapter, we explore the different patterns of development following the hatching of the crustacean larvae. For many groups of crustaceans, the free-​living, postembryonic, and prejuvenile phase is by far the most important part of their life cycle, providing the link between different life modes in successive phases (e.g., between a sessile adult life and the need for long-​range planktonic dispersal). Among the aspects covered, we discuss the specific criteria for what a “larva” is, including the necessity for defining specific larval traits that are lacking in other phases of the life cycle. We examine the typical anamorphic and hemianamorphic developmental patterns based on larval examples from a wide selection of groups from Decapoda to Copepoda, Thecostraca to Branchiopoda. In these groups, we examine the most common larval development patterns (including intraspecific variability) of, for example, the zoea, furcilia, copepodite, nauplius, and cypris larvae. We also expand on the importance of the molting cycle as the main driver in larval ontogeny and evolution. Finally, we discuss some of the more general trends of crustacean larval development in light of the general patterns and latest knowledge on tetraconate and arthropod evolution.

INTRODUCTION Like some vertebrates (fish, amphibians) and many invertebrates (e.g., mollusks, polychaetes, insects, echinoderms), most Crustacea pass through a complex life cycle that comprises an embryonic phase inside the egg membrane, posthatching development through a larval phase, and the juvenile–​adult phase. Complex life histories, especially in marine species, are often referred to as biphasic, which refers to two contrasting types of environment in which the different life history stages are typically found: (1) the juvenile, adult, and embryonic parts of the life cycle are passed Developmental Biology and Larval Ecology. Edited by Klaus Anger, Steffen Harzsch, and Martin Thiel. © 2020 Oxford University Press. Published 2020 by Oxford University Press.

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Developmental Biology and Larval Ecology in the benthos (on the bottom of aquatic environments) or on land; (2) larval development occurs mostly in the plankton (freely floating in the water column). Larvae differ from conspecific juveniles and adults based on morphological, ecological, behavioral, physiological, and/​or other biological traits. In most marine Decapoda and other aquatic crustaceans, the juveniles and adults are benthic deposit feeders or predators, whereas the larvae develop in the pelagic environment, feeding on other plankters. Other crustaceans also have actively swimming planktonic larvae except for sessile (e.g., Cirripedia, Thoracica) or parasitic juveniles and adults (e.g., Rhizocephala, caligid Copepoda) (Figs. 6.1-6.3). Another example of ontogenetic changes in lifestyle is found among taxa in which the juveniles and adults live in fresh water or on land, whereas their larvae survive only in physically stable marine environments [larval “export” strategies (Anger et al. 2015)], and others have an abbreviated larval phase or direct development. This chapter gives a brief overview of the most common larval forms and developmental patterns in the Crustacea. In view of the comprehensive and recently published Atlas of Crustacean Larvae [henceforth referred to as ACL (Martin et al. 2014a)], a new review has become a major challenge. In the ACL, the morphology of virtually all known types of larvae from numerous taxa is exhaustively reviewed, including both extant and fossil forms, as well as embryonic “larval” stages found only inside the egg membrane (e.g., Peracarida). Here, we largely exclude fossil larval forms as well as those recognized only during embryogenesis (not free living). This chapter bridges the preceding parts of this volume, dealing with embryonic development and hatching, with the subsequent chapters addressing larval ecology, growth, feeding, dispersal, and other aspects. Both the scope and the potential audience of this chapter differ from that of the ACL. The latter provides an almost complete reference volume for researchers specialized in taxonomy, larval biology, morphology, or life history evolution of the Crustacea. This chapter, in contrast, offers a brief introduction for (1) students interested in crustaceans and their larvae; (2) researchers working on crustacean ecology, physiology, and so forth, with an interest in the ontogeny of those aspects; and (3) comparative larval biologists studying taxa other than crustaceans.

THE FUNDAMENTAL QUESTION: WHAT MAKES A LARVA AND WHAT DOES NOT? In the ACL, the term larva is broadly defined as “any immature, post-​embryonic form of an animal that differs morphologically from the adult and often develops into the adult either gradually (via anamorphosis) or by more abrupt changes in morphology (metamorphosis)” (Martin et al. 2014a, p. 2). Although this definition is correct and normally useful, the question remains to what extent do immature forms must “morphologically differ” from the adults to qualify them as “larvae?” In young brachyuran crabs, for instance, the carapace is often initially longer than it is broad, whereas adult crabs typically show opposite proportions (Bolla et al. 2014). Thus, at least minor morphological differences can also occur between juveniles and adults. Can we always distinguish unambiguously between larvae and juveniles, and which are the decisive criteria? Absence of Adult Features Among the morphological disparities between larvae and conspecific adults, a lack of adult features is considered to be a significant characteristic of a larva. However, juveniles also lack some adult traits. Primary sexual organs appear only when maturity is reached, secondary sex-​specific characters are not present at hatching, and some adult appendages may initially be absent or nonfunctional (Negreiros-​Fransozo et al. 2007, Bolla et al. 2014). In crayfish, the hatchling is nonfeeding and may remain attached to the egg case, and the uropods and some feeding and locomotory appendages



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(thoracopods, pleopods) as well as several sense organs (e.g., various aesthetascs) are typically absent, incomplete, or not yet functional (Vogt 2013). These adult traits are developed stepwise during later molts. Despite morphological and functional incompleteness, these early posthatching stages are considered as juveniles with “embryoid” features (Minelli and Fusco 2013) and not as larval stages. Therefore, it is generally accepted that all crayfish have a direct development (Goy 2014). As in crayfish, similar traits have also been observed in several species of caridean shrimp with “larval development” that reportedly consists of a single stage. This pattern, commonly referred to as completely or extremely abbreviated, is known for some hololimnetic palaemonids and atyids (Magalhães 1988, Jalihal et al. 1993). In these species, the newly hatched “larva” (mostly referred to as zoea) lacks uropods and is typically nonfeeding, but otherwise similar to an adult, equipped with functional pereopods, segmented antennae, chelae, and pleopods. These traits allow for a free-​living adult-​like lifestyle, with benthic walking and swimming immediately from hatching. Hence, these early postembryonic stages are morphologically and behaviorally even more advanced (i.e., more similar to the adults) compared to the early posthatching stages of crayfish. The initially underdeveloped characteristics appear during the following one to three molts (Rodriguez and Cuesta 2011). We thus suggest that such early postembryonic stages of species with an “extremely abbreviated larval development” should be considered morphologically incomplete juveniles rather than larvae (i.e., such species show a direct development). Similar to crayfish, the initial stages may still show some remaining larval traits such as nonfeeding behavior and an absence of functional walking or feeding appendages. In conclusion, a distinction between larvae and juveniles based solely on morphological or functional incompleteness is ambiguous, and will vary among taxa and authors. We therefore propose that a mere absence of adult features is a sine qua non; but, as with any argument based solely on the absence of evidence, it is not a sufficient precondition for calling an early postembryonic stage a larva. Presence of Specific Larval Features Larvae showing ontogenetically specific adaptations and lifestyles are known from insects (e.g., lepidopteran caterpillars, aquatic dragonfly, and mayfly larvae) and amphibians (e.g., tadpoles). Further examples are found in the planula (coelenterates), the trochophore (entoprocts, mollusks, annelids, echiurans, sipunculans, nemerteans), the veliger (gastropods, bivalves, scaphopods), and the pluteus (echinoderms). Larval forms can be so different from the conspecific adults that some of them have been described as distinct species, and many larval names were originally genus names. Among the Crustacea, this applies, for example, to the nauplius, cypris, calyptopis, furcilia, amphion, zoea, megalopa, nisto, and phyllosoma. Differential adaptive morphologies in early and late larval stages versus conspecific juveniles or adults are illustrated in Figure 6.1. Among the examples shown, the most conspicuous ontogenetic disparities in morphology and lifestyle can be seen in barnacles (Cirripedia, Thoracica) (Fig. 6.1A–​C) (Chan et al. 2014). The adults are permanently attached to benthic substrates or other solid surfaces such as rocks, whale bodies, mollusk shells, or large decapods, whereas the hatching stage is a nonfeeding planktonic nauplius larva (Figs. 6. A–​C, 6.3B–​G, and 6.4A) (Martin et al. 2014b). After passing through a planktivorous phase with several naupliar molts in the water column, a brief, nonfeeding cypris stage is reached. Nauplii and cypris larvae differ so much morphologically, behaviorally, and ecologically from the sessile juveniles and adults, that their uniqueness within the life cycle identifies them as true larval stages (Fig. 6.1A–​C, but compare with Figs. 6.3B–​G and 6.4A). Striking differences between larvae and adults are also typical of spiny and slipper lobsters [Achelata (Palero et  al. 2014)]. Their hatching stage is characteristic of phyllosoma larva

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(Fig.  6.1  D). The subsequent planktonic development is very extended, comprising numerous molts and lasting several months or even greater than one year. The last phyllosoma stage molts to a morphologically quite different form called puerulus in the Palinuridae or nisto in Scyllaridae (Fig. 6.1E). Behaviorally comparable to the cypris of the cirripedes, this stage is freely swimming and, at least in some taxa, nonfeeding, searching for a benthic habitat suitable for settlement. Morphologically, however, this stage is already an adult-​like juvenile. A  combination of juvenile morphology with larval behavior is found also during postembryonic stage IV of nephropid (clawed) lobsters. Their epibenthic searching and swimming behavior disappears only gradually through one or more molts before settlement occurs and fully benthic crawling begins (Fig. 6.1F) (Charmantier et al. 1991, Goy 2014). We propose that such freely swimming but otherwise adult-​ like settling stages of achelate and clawed lobsters should be considered morphologically incomplete juveniles with some remaining larval traits in locomotor (and maybe feeding) behavior, but not larvae. These examples show that typical (or true) larvae show a lifestyle that may differ substantially from that of conspecific adults, and this requires specific transitory adaptations in morphology, behavior, and physiology. We therefore propose that larvae are primarily characterized by the presence of larval features rather than a mere absence of adult traits. In crustaceans, typical larval traits occur most obviously in the functional morphology of locomotor and feeding appendages (Williamson 1969, Williamson and Bliss 1982, Anger 2001). These disappear, or are transformed, at the end of the larval phase and are thus absent in the subsequent juvenile and adult life history stages. However, in some taxa with gradual (or anamorphic) patterns of postembryonic development (e.g., many Caridea, Euphausiacea, Branchiura; discussed later), functionally insignificant vestiges of larval traits may persist into the initial juvenile phase while all adult traits (except for sexual organs) are already present. In these cases, the presence of typically adult features, in combination with a loss of functionality of larval traits, is not sufficient to define such stages as larvae. Instead, we may refer to them as “early juveniles with gradually reduced vestiges of larval features.” This implies that gradual developmental patterns may not have a clear point of transition (i.e., one specific molting event) from the larval to the juvenile phase.

DEFINING CONCEPTS AND LEVELS OF DEVELOPMENT The course of development can be divided into morphologically or otherwise (e.g., ecologically, behaviorally) distinguishable periods, which are commonly referred to as stages. This term is unspecific and is used inconsistently, denoting any distinct developmental point in time, and it may be applied at various ontogenetic levels. This includes the distinction of different life history stages (embryonic, larval, juvenile, adult), larval stages (e.g., nauplius I, nauplius II, cypris), or molt-​cycle stages (recurrent integumental changes occurring between successive ecdyses; described later). When there is confusion between the different meanings of the term stage, it is necessary to add determiners such as life history, larval, or molt cycle.

Fig. 6.1. Larval forms in crustaceans. Comparison with adults (examples). (A–​C) Cirripedia: Tetraclita squamosa and (C) T. formosana. (A) Nauplius. (B) Cypris. (C) Sessile adult. (D–​F) Achelata (Palinuridae). (D) Palinustus sp. phyllosoma. (E) Palinustus sp. puerulus. (F) Palinurus elephas adult. (G–​I) Brachyura: Dyspanopeus sayi. (G) Zoea. (H) Megalopa. (I) Adult. ( J–​L) Caridea: Palaemon zariquieyi early ( J) and late (K) zoea. (L) Adult. (A–​C) Photos courtesy of B. Chan (D) Photo courtesy of G. Guerao. (E) Photo courtesy of CSIRO. (F–​I) Photos courtesy of G. Guerao. ( J, K) Photos courtesy of K. Anger. (L) Photo courtesy of B.A. Galán. See color version of this figure in the centerfold.

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Developmental Biology and Larval Ecology Similar stages can be assigned collectively to a particular ontogenetic phase. Like stage, however, phase may also be used at different levels of development. On the life history level, these two concepts are used as synonyms (e.g., embryonic life history stage = embryonic phase). Within the larval phase of the life cycle, a sequence of morphologically similar stages, such as the naupliar stages of cirripedes or the zoeal stages of decapods, can again be joined in a particular (e.g., naupliar, zoeal) phase. Successive stages belonging to the same phase are sequentially numbered (nauplius I, II; zoea I, II, etc.) (see, for example, Figs. 6.3A, 6.5, and 6.6). The Fundamental Process in Arthropod Ontogeny: The Molting Cycle In Arthropoda, including crustacean larvae, the process of molting subdivides the development and growth into distinct periods. Within individual larval stages such as zoea I or II, we can distinguish successive stages in the cuticular and epidermal morphology. These cannot be identified normally as changes in external morphology or body size (exceptions are described later), but as microscopically visible modifications in the histology of the integument. Drach’s classification system is used to describe these changes (Drach 1939). In contrast to juvenile and adult crustaceans, for which such periodic anatomical alterations as well as their underlying hormonal control mechanisms are known in detail, there have been few studies on molting cycles among larvae. Such knowledge would shed more light on larval physiology, ecology, and behavior, all of which may vary greatly within single molting cycles. For instance, a single zoeal stage of a crab can show up to a threefold increase in organic biomass as well as great variations in hormone titers, rates of feeding, respiration, and ammonia excretion, whereas its external morphology and body size remain unchanged (Anger 2001). Obviously, many internal organ systems develop gradually through successive molting cycles. This has been shown for the nervous system of Branchiopoda (e.g., Triops cancriformis seen in Fig.  6.2C) (Fritsch and Richter 2010)  and the cement gland of rhizocephalan cypris larvae (Fig. 6.4A) (Høeg and Ritchie 1987). The potential for intrastage growth and changes in external morphology is also considerable in several groups. This ability has seemingly evolved several times, particularly in some of the parasitic groups in Thecostraca and Copepoda. Nonmolting growth, not only in the size of the genital segment, but also in whole-​body size, is well known from siphonostomatoids such as the pennelid Phrixocephalus (Boxshall 1986). This was not thought to be the case for larvae, but Eichner et al. (2015) showed recently that larval growth of salmon louse (Lepeophtheirus salmonis) is considerable (Fig. 6.3A). This fact may explain why morphometric data cannot be used easily as valid stage indicators. In L. salmonis, the correct number of chalimus stages was only discovered using a large data set and a statistical approach (Hamre et al. 2013). Another well-​known example of nonmolting (intrastage) growth is the extreme growth of the vermigon in the Rhizocephala. This stage grows from a worm-​like appearance (ca. 100–​150 µm long), into the fully extended rootlet system (interna), and later the egg sac (externa), which can be several centimeters in length (Fig. 6.4) (Bresciani et al. 2001, Glenner 2001). The dependence on external food sources (relevant for cultivation) varies dramatically during the molting cycle (Anger 2001). In planktivorous decapod larvae, the early stages in the cycle (postmolt and intermolt) are characterized by high rates of feeding and biomass accumulation. Fig. 6.2. Anamorphic developmental patterns. (A–​D) Examples of free-​living Branchiopod early larvae showing anamorphic development. (A)  Rehbachiella kinnekullensis (fossil). (B)  Artemia salina. (C)  Triops cancriformis. (D) Lynceus brachyurus. (E–​H) Scanning electron micrographs of the first four larval stages of Limnadopsis parvispinus (Branchiopoda, Spinicaudata), an Australian clam shrimp. (A–​D) Modified from Olesen (2003), with permission from Taylor and Francis. (E–​H) Modified from Pabst and Richter (2004), with permission from Elsevier.

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Developmental Biology and Larval Ecology After passing through a critical point in the cycle, the D0 threshold, all subsequent (premolt) stages, become nutritionally autonomous. Post D0, a larva can develop successfully through the remaining 50% to 70% of the molting cycle, reaching the next larval stage even in the complete absence of food (Anger 1987). Likewise, patterns of larval migration behavior and response to environmental cues (e.g., light, gravity) may change during successive stages of the molting cycle, with implications for larval dispersal in the plankton, recruitment, genetic connectivity, and so on (review: Anger et al. 2015; see also Chapters 12 and 14 in this volume). Stage versus Instar Because larval crustaceans generally show a stepwise increase in the complexity of locomotor and feeding appendages, usually also in body size after molting, ecdyses are commonly used to define successive larval stages. In some cases, however, molting is not associated with significant morphological changes, and sometimes not even with an increase in body size (mark-​time molting) (Gore and Schram 1985). Gradual patterns of development with little or no morphological changes at ecdysis are typically found in juvenile growth, but occur also in larvae, especially in those of euphausids (Makarov 1974, Feinberg et al. 2006) and caridean shrimp (Walsh 1993). In such cases, the neutral term instar rather than stage is often used to denote successive molt cycles. It is independent of the phase of the life cycle and the presence or absence of morphological changes (Anger 2001). Successive instars may thus be morphologically identical (e.g., in juveniles), similar (in successive zoeal stages), or completely different from each other (e.g., final-​stage crab zoea vs. megalopa; Fig. 6.1). The terms instar and stage are often used as synonyms, although the latter generally implies at least some morphological change whereas the former does not. The unclear use of developmental terms is confusing but likely unavoidable as a result of differing views of different authors and research schools, and the numerous taxa and developmental patterns found in Crustacea. This terminological diversity therefore requires that each author clarify unambiguously what is actually meant. Because a specific ontology-​based terminology covering evolution and development is not yet available for Crustacea [but work is underway; see, for example, Vogt et al. (2012)], clarity can be achieved by using specific determiners (as mentioned earlier) and adding clarifying information, as well as refraining from inventing new terminology. This is particularly important in cases with gradual (anamorphic) developmental transitions that do not permit an application of a strict classification system.

PRINCIPAL TYPES OF CRUSTACEAN LARVAE Nauplius (pl. nauplii) The nauplius larva is probably the most widespread larval form in the Crustacea. Originally, the term was used to describe the first larval stage of a copepod, but it is now used in a more general way for various groups of crustaceans. The ACL gives an authoritative account on almost every aspect of nauplii (Martin et al. 2014b). A typical nauplius consists of only three fifths of an adult crustacean head, and it is found in practically every marine environment. If present, it is always the earliest larval stage, and it is generally believed that most recent Crustacea pass through a naupliar phase during their ontogeny, at least during embryogenesis. Several major groups have free-​swimming nauplii in early development, albeit with differences in morphology. The orthonauplius has only three pairs of appendages: the first and second antennae and the mandibles (typically carrying long mandibular palps and natatory exopods), showing no further segmentation or limb anlagen. The best examples of “true” orthonauplii can be found in the Branchiopoda (e.g., Artemia), Copepoda [many Harpacticidae (Dahms et  al. 2007), Ostracoda, and Remipedia (Dahms et  al. 2006)]

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Fig. 6.3. Hemianamorphic [metamorphic sensu Atlas of Crustacean Larvae (Martin et al. 2014a)] development patterns exemplified (early anamorphic phase followed by metamorphic change). (A)  Developmental stages of Lepeophtheirus  salmonis from copepodite to young adult. Notice the strong potential for intrastage growth shown, for example, in the chalimus 2 stage. (B–​G) Nauplius stages 1 through 6 of Balanus improvisus. cs, caudal spine; hs, head shield spine; la, labrum; ms, median spine; s1, first pair of spines; s2, second pair of spines; s3, third pair of spines. (A) From Eichner et al. (2015), with permission from Elsevier. (B–​G) Modified from Semmler et al. (2009), with permission from Elsevier. See color version of Fig. 6.3A in the centerfold.

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Developmental Biology and Larval Ecology (described later) ( Figs. 6.1A, 6.2, 6.3B–​G, and 6.4A, B, D, E). All nauplii of Thecostraca have characteristic frontolateral horns (Figs 6.1A and 6.3B–​G) (Høeg et al. 2009). When additional limb primordia and somites are present, the term metanauplius is used (Fig. 6.2). These larvae typically have a more complex mouth apparatus with a labrum. A metanauplius can follow an orthonauplius, as seen, for example, in Artemia (Fig. 6.2B) (Cohen et al. 1999) or some Copepoda; but, very importantly, it can also be the hatching stage as in Cephalocarida (Sanders 1963). The hatching stage of the common carp louse Argulus foliaceus, which has a full complement of head appendages and thoracopod anlagen, is also a metanauplius because it retains specific larval traits, such as the naupliar swimming apparatus (Møller and Olesen 2014). Many of the malacostracan groups (especially Decapoda) pass the nauplius stage within the egg (egg–​nauplius); the Dendrobranchiata and Euphausiacea are exceptions with a free-​living (but lecithotrophic) nauplius. The euphausiacean nauplius passes through two stages. In broadcast-​ spawning species, which shed their eggs directly into the water column, the second nauplius molts directly to a metanauplius. In sac-​spawning species (embryos develop in a membranous sac on the last thoracopods), metanauplius stages are passed inside the brood pouch; some authors use the term pseudo-​metanauplius for the early stage, but this term is discouraged (Martin and Gómez-​ Gutiérrez 2014). All these stages are nonfeeding. It has been suggested that the nauplii of Dendrobranchiata and Euphausiacea were evolutionary novelties of secondary origin based on their lack of a feeding apparatus (Scholtz 2000). This hypothesis was recently supported by Jirikowski et al. (2015), who found differences in the developmental genetic control of muscle development. However, the malacostracan ground pattern was not reconstructed, and thus no conclusion on the absence or presence of nauplii in the ground pattern of this major group could be made. In another recent article, Akther et al. (2015) presented morphological evidence speaking against the Scholtz hypothesis. By using an outgroup comparison method, they argued convincingly for nauplii being ancestral to the Malacostraca and subsequently lost (or heavily modified) in several groups. Calyptopis and Furcilia In the Euphausiacea, the naupliar phase is followed by a larval form termed calyptopis (first feeding stage). Some traits of a late metanauplius persist in three successive stages, such as natatory antennae, but also zoeal traits are found. In the thoracic appendages, only the maxillipeds are functional whereas the pereopods and pleopods appear only as limb buds. The larvae generally look shrimp-​like, with the carapace and a well-​defined telson appearing in the first stage, and pleomeres and uropods in the second and third stage. The three calyptopis stages are followed by several furcilia stages. There is no metamorphic molt from a late furcilia to an early juvenile, and thus this specific larval morphology is disappearing gradually. This is evident in the development of the pereopods, pleopods, and antennae, all of which gradually lose their natatory function. Cypris The cypris larva is found exclusively in the Cirripedia, the major group to which the sessile barnacles and parasitic Rhizocephala belong. This larval stage is named after a genus of Ostracoda (Podocopida: Cypris) because they were mistaken for adults of this group (Newman 2005). It is the final, nonfeeding, free-​living stage, showing a characteristic morphology. The bilobed carapace (developed from the head shield of the nauplius) contains the chemosensitive lattice organs (Høeg et al. 1998) and covers a body with six natatory thoracopods. In all Cirripedia, the first antennae are prehensile, unique among Crustacea, and highly modified for “exploratory walking” on the substratum before settlement. The nauplius eye persists, but also a complex eye is present (occurring



Patterns of Larval Development

exclusively in this life cycle stage of Thoracica) (Lagersson and Høeg 2002, Blomsterberg et  al. 2004). The cypris is transitional between the pelagic larval and the juvenile/​adult phase, changing to a filter-​feeding or parasitic life. The importance of both phases is seen in the highly specialized cyprid morphology featuring highly complex first antennae and six pairs of thoracopods that allow for efficient swimming. The settlement decision made by the cyprids is crucial for the survival and reproduction of the sessile adults (Høeg and Møller 2006; see also Chapter 8 in this volume). The third antennal segment is equipped with a flattened structure that is densely covered with minute setulae; and the fourth (distal-​most), with highly specialized chemo-​and mechanosensory setae. Also, the highly complex muscle and tendon system of the antennae enables it to perform very intricate and advanced movements and turns during substratum exploration (Fig. 6.5A, 1 and 2) (Bielecki et al. 2009, Maruzzo et al. 2011). The walking phase ends with the cyprid selecting a spot for the final life phase, either as sedentary (selective) filtrator in most barnacles, or penetrating into a host in the Rhizocephala (described later; vermigon and kentrogon) (Fig. 6.5). The cypris generally follows the last nauplius stage, but a number of species hatch as cyprids (e.g., some Scalpellum species, all akentrogonid Rhizocephala) (Crisp and Knight-​Jones 1953, Høeg 1995, Kolbasov et al. 2008). In general, cypris larvae are of quite uniform morphology across the whole Thoracica and Thecostraca. Kentrogon and Vermigon These specialized and highly derived larval stages are found in one of the two major groups of parasitic Rhizocephala: the Kentrogonida. Only a few species have been described in detail, but the presence of specialized larvae is probably one of the apomorphic traits of the group. After settling on the surface of the future host (typically in the mantle cavity of a decapod), the cypris larva undergoes a metamorphosis into a kentrogon larva. This stage is completely enclosed by the cypris cuticle at first, but as the metamorphosis completes during approximately 48 hours (including, for example, complete loss of the thoracopods), the distinctive kentrogon larva is formed and (in most species studied) the cypris cuticle is shed. Like the cyprids, the kentrogon remains cemented to the host with the first antenna and undergoes another (internal) metamorphosis to the so-​called vermigon stage (Fig. 6.5C). This worm-​like stage is injected by the kentrogon into the host via a stylet formed between the antennae by the kentrogon and is the most highly derived larval form found in the Crustacea. It is covered by a thin cuticle and has internal structures with specialized cells. The vermigon migrates through the body of the host and develops into the adult interna/​ externa (Høeg and Lützen 1995, Glenner et al. 2000, Glenner 2001, Walker 2001, Høeg et al. 2012). Trichogon Kentrogon/​vermigon larvae only develop from those cyprids destined to become female adults, whereas another larva (or larval-​like stage) develops from the cyprids destined to be males: the so-​called trichogon. This larva is vermiform and covered in a spiny cuticle, and it develops within the late cypris larva. It carries the precursors to spermatogonia and, following the injection of the stage into the female mantle cavity, it molts without reforming a cuticle and transforms to a cuticle-​ free dwarf male that produces sperm (Høeg 1987, Glenner and Høeg 1994, Høeg and Lützen 1995). Copepodite The copepodite larva is limited to the Copepoda, where it is ubiquitous in both parasitic and free-​ living groups. Similar to the nauplius phases of Cirripedia, Copepoda, and many other groups, the copepodite phase contains three to five separate stages (Fig. 6.5C). Again, similar to nauplii, this

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Developmental Biology and Larval Ecology number is reduced in parasitic forms, with typically only two copepodite stages (see Fig. 6.5A). The copepodite always follows the nauplius phase, after a metamorphic molt from the last nauplius or metanauplius. The first copepodite thus has an advanced morphology compared to the nauplius, with all five cephalic appendages developed. Huge first antennae develop into long, multiannulated appendages typical of many adult Copepoda. Three thoracic appendages are developed, the first one as a maxilliped. The abdomen (sometimes called a urosome in Copepoda) and telson also appear, giving the copepodite almost a “small adult” look—​hence its name. During the copepodite phase, more segments are added at each molt, and significant growth of especially the thorax takes place, although there is considerable variation among the major lineages (Ferrari et al. 2010, Martin et al. 2014a). The last molt from a copepodite to a juvenile is an anamorphic molt, as the last copepodite closely resembles the adults (Fig. 6.5C). In the parasitic groups, the copepodite is generally the infective stage, seeking out, for example, the host fish in caligid Copepoda (Dojiri and Ho 2013). During the last copepodite stage, a thread-​like, medial–​anterior outgrowth is everted (first visible coiled up inside a pouch during the early stage) and used to tether the larva to the host fish (Figs. 6.3A and 6.5A). It is erroneously called a frontal filament in Copepoda literature. The correct use of frontal filament is disputed, but it should be limited to the specialized frontal sensory organ found in Cirripedia. This attachment serves the larva through the molt to the chalimus, and through all the stages of this phase. There has been considerable interest from marine aquaculture research groups into how precisely the copepodites find their hosts, as few species of caligids (e.g., Caligus rogercresseyi, C. elongatus, Lepeophtheirus salmonis) cause severe damage to the farmed salmon industry (Tully and Nolan 2002, Boxaspen 2006, Mordue Luntz and Birkett 2009, Torrissen et al. 2013). Chalimus The chalimus larva is only found in parasitic Copepoda of the Siphonostomatoida, being widespread and of high commercial interest. Instead of molting from the last copepodite to the adult, this characteristic and specialized larval form is intercalated. It is attached to the host fish by a thread-​ like anterior outgrowth and, as the larva molts, a bulbous growth forms at the base of the filament and the larval cuticle is shed only on the proximal side of this. So for each of five or six molts, a new part is added proximally. The widespread salmon louse L. salmonis only has two chalimus stages and produces a new filament for each (Fig. 6.3A) (Piasecki and Mackinnon 1993, Bron et al. 2000, Gonzalez-​Alanis et al. 2001). Successive chalimus stages increase in size and number of segments to reach the adult condition. Although the attached copepodite might start to feed, the main host damage occurs in the chalimus stages, when the mouth tube is almost of adult shape, allowing for increased feeding ( Jones et al. 1990). The other appendages used for attachment to the fish as adults are also developing during the chalimus phase (e.g., second antennae and the specialized cephalothoracic suction cup) (Anstensrud 1990, Boxshall 1990). There is considerable confusion about the phase following the chalimus. Some authors identify up to two preadult stages whereas others fail to differentiate these stages from the adult (e.g., Tully and Nolan 2002, Burka et al. 2011, Martin et al. 2014a). Isopod Larval Stages Zuphea/​Praniza The manca stage of peracarid Isopoda is difficult to include among the true larvae and is only recognizably different from the adults by details of the posterior thorax (lacking the last thoracopod). However, because some of them have a highly derived semi-​or fully parasitic lifestyle, which differs substantially from that of the adults, we include some examples here. An example of a manca stage is the zuphea/​praniza larva found exclusively in gnathiid isopods. It is the only life history stage that



Patterns of Larval Development

feeds in this group (highly unusual in crustaceans), and it is one of the very few known examples of truly parasitic larvae in the Crustacea. It occurs in different habitats, including rock pools, estuaries, and coral reefs, where it can cause damage to their hosts [teleost and elasmobranch fish (Smit and Davies 2004)]. The two names zuphea and praniza cover the same developmental stage. Zuphea is used for unfed larvae whereas praniza is used for larvae replete with host blood. Although only a few species are known in detail, it is generally known for most common species that the zuphea I is released from the female marsupium and takes its first blood meal turning it (per gnathiid traditional nomenclature) to a praniza I (recognizable by the considerable swelling of the thorax). After a period of roughly five to eight days, it molts to the zuphea II. This cycle is repeated until the praniza III molts into the adult (Smit and Davies 2004, Wilson et al. 2011). In the parasitic isopod groups Bopyridae and Cryptoniscidae, the life cycle is completed exclusively on crustacean hosts. It includes two pelagic (infectious) larval stages (epicaridium and cryptoniscus) and two parasitic on-​host stages (microniscus and bopyridium) (Williams and Boyko 2012). The epicaridium hatches and seeks out a copepod (typically a calanoid) intermediate host, on which it metamorphoses into the microniscus, which grows considerably in size (up to 10 times) while on the host. The microniscus leaves the host and transforms to the cryptoniscus, seemingly without molting, but only by expanding folds in the cuticle. This process again underscores the peculiarities of isopod larvae in general and questions the validity of this particular stage, although intrastage growth is known from other groups of Crustacea (as mentioned earlier). The expanded cryptoniscus then seeks out the final host, typically a crab, on which it settles between the gill lamellae. The settled larvae molt to the juvenile, which is also termed bopyridium by some authors (Anderson and Dale 1981, Boxshall 2005, Williams and Boyko 2012). Zoea The zoea is the typical taxonomically most widespread planktonic larval form of the Decapoda (Williamson 1969, Williamson and Bliss 1982). An equivalent form, the “amphion” larva occurs in the little-​known eucarid taxon Amphionidacea, which is considered to be the possible sister taxon of the Decapoda (Kutschera et  al. 2012). A  zoea is characterized by the presence of natatory exopods on some or all thoracopods. At hatching, the number of natatory maxillipeds varies among taxa between one and three. The amphion hatches with only the maxillipeds developed for swimming. Throughout their zoeal phase, the Anomura and Brachyura exclusively use the first and second maxillipeds for swimming, whereas all other decapod groups are comprised of at least some taxa that also use the third maxilliped (Fig. 6.6A). The number of maxillipeds and pereopods with natatory exopods expressed at hatching vary among taxa. Although the Amphionidacea, Caridea, Thalassinidea, Anomura, and Brachyura lack natatory thoracopods, the Stenopodidea, Polychelida, and Achelata hatch with one to three; and the Astacidea, with five (Ingle 1992). In Caridea, the late zoeal stages have functional pereopods, whereas in Brachyura and Anomura, the third maxillipeds and pereopods appear only during the final zoeal stage as limb primordia without setae. Pleopods remain absent or rudimentary and nonfunctional (Fig. 6.6B). Except for the Dendrobranchiata, the Decapoda (all Pleocyemata) hatch as zoeae. The Dendrobranchiata hatch as a nauplii (like the Euphausiacea) and pass subsequently through a naupliar phase comprising of five to six stages, followed by a zoeal phase with taxon-​specific numbers of stages (Martin et al. 2014b). In decapods with abbreviated larval development (described later), hatching occurs as a morphologically advanced zoeal form, commonly showing nonfunctional pleopod primordia. In extreme cases, the entire zoeal phase is skipped and hatching occurs as a megalopa or even as a juvenile (direct development).

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Fig. 6.4. (A) Major events in the metamorphosis of pedunculated (1–​4), sessile (5–​10), and parasitic (11–​15) cirripedes. (1–​5) Lepas; (1) cyprid on substratum by the antennules, (2) whole cypris body raised on attachment point with developing shell plates visible, (3) last cypris molt with two variations in Lepas (3a) and L. anserifera (3b), (4) juvenile starts feeding. (5–​10) Megabalanus rosa; (5) cyprid on substratum by the antennules, (6) contraction of antennular muscles pulls the body close to the substratum, (7) whole cypris body raised on the attachment point, (8) last cypris molt, (9) juvenile is still flexible and there are still no signs of shell plates, (10) juvenile starts feeding and developing shell plates. (11–​15) Sacculina carcini; (11) cyprid cemented to substratum (base of crab’s seta) by the antennules; (12) after a metamorphic molt, the kentrogon, which remains within the cypris carapace, is formed; (13) after a second molt a new kentrogon (with a stylet) is formed; (14) the stylet penetrates down the setal canal and the vermigon is injected (also comprises a molt); (15) the vermigon breaks free from the tip of the stylet and flows into the host’s hemocoel. (B, C) Light microscopic images of the process from figure D13–​D15. (B) Kentrogon with formed injection stylet on crab seta (corresponds to D13). (C) Injected vermigon is visible to left in the picture (would have been inside the crab coelom under natural conditions) and corresponds to D14 and D15. Modified from Høeg et al. (2012), with permission from Oxford University Press. See color version of this figure in the centerfold.

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Fig. 6.5. (A) Life cycle of Lepeophtheirus salmonis (early anamorphic phase followed by metamorphic change). Note the presence of anamorphic as well as metamorphic molts. (B, C) Six nauplius stages (N1– N6) of Eudiaptomus vulgaris (calanid copepod) are followed in (C) by five copepodite stages (C1– C5). Note that the nauplius stage molts are anamorphic whereas the N6 through C1 molt is metamorphic. (D E) Lepeophtheirus salmonis, nauplius stages 1 (D) and 2 (E). (A) From Piasecki and Avenant- Oldewage (2008), with permission from Taylor and Francis. (B, C) Modified from Dussart and Defaye (2001), with permission from John Wiley and Sons. (D, E) Modified from Schram (2004), with permis­sion from Cambridge University Press.

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Developmental Biology and Larval Ecology Zoea larvae from different decapod taxa are referred to by various names, reflecting morphological variation among taxa but also different terminological traditions. In the Dendrobranchiata, the early zoeal stages are called protozoeae. In contrast to the typical zoeae of the Pleocyemata, all cephalic appendages are fully developed, and the first antennae are natatory (as in a nauplius). These protozoeae are followed by several mysis stages, which are typical zoeae resembling those of the Caridea. Other names for zoeal forms are the eryoneicus of the Polychelida as well as the naupliosoma and the phyllosoma of the Achelata (Palero et al. 2014). A naupliosoma is comparable to the prezoea that may occur in all decapod groups; it is generally considered a prematurely hatched embryonic stage of very short duration, rather than a true larval stage (Williamson and Bliss 1982, Anger 2001). Decapodid Kaestner (1980) proposed the name decapodid to denote the final larval phase of the Decapoda preceding the first juvenile instar. It is characterized by an appearance of uropods as well as functional pleopods bearing long, plumose natatory setae; the pereopods are walking legs, with or without exopods (Williamson and Bliss 1982). These changes in functional morphology allow for

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Fig. 6.6. Patterns of postembryonic development in malacostraca Decapoda. (A) Hemianamorphic pattern in Brachyura (Chiromantes ortmani). (B)  Anamorphic patterns in Caridea (Palaemonetes argentinus). Pattern fills of appendages: black, functional appendage; white, bud or nonfunctional appendage; dotted, reduced. Numeration of the appendages (1–​19):  antennule, antenna, mandible, maxillule, maxilla, maxillipeds 1–​3, pereopods 1–​5, pleopods 1–​6 (pleopod 6 = uropod). M, metamorphosis; P, pleonite; T, telson; Z, zoea. Modified after Magiera (2009) and Guerao et al. (2012).



Patterns of Larval Development

a combination of benthic crawling and demersal swimming. The decapodid is thus transitional between a pelagic zoea and a benthic juvenile, and it differs clearly from both of these phases. This phase generally consists of a single stage. It is mostly referred to as a megalopa (Fig. 6.1H) [Brachyura; Anomura; Gebiidea and Axiidea (=Thalassinidea)]. In the Anomura, the name glaucothoe is also frequently used. In Polychelida, the equivalent stage is called eryoneicus (Martin 2014). In the Achelata, the first postzoeal stage (commonly named puerulus in the Palinuridae, nisto in the Scyllararidae) shows a behavioral transition from pelagic swimming to benthic crawling. Although this developmental stage is completely different from the preceding phyllosoma stages, only minor morphological differences from the subsequent juveniles have been found, such as a “somewhat dorsoventrally flattened” body shape and “proportionally large natatory pleopods” (Palero et al. 2014). This stage is probably nonfeeding, searching for a suitable benthic settlement habitat. An ontogenetically transitory independence from food in a settling stage also occurs in the megalopa of some hermit crabs and in the cypris stage of barnacles [termed secondary lecithotrophy (Anger 1989)]. This nutritional independence relies on energy reserves accumulated during the preceding zoeal (or naupliar) phase, in contrast to primary lecithotrophy subsequent to hatching, which is based on an enhanced female energy investment in egg production per offspring. The nonfeeding puerulus and nisto stages have nonfunctional mouthparts (mandibles, maxillae), resembling those of the early postembryonic stages of crayfish (Vogt 2013). The hermit crab megalopa and the cypris, in contrast, not only lack some adult characteristics (benthic feeding) but also have natatory thoracopods (i.e., larval traits). These disappear after metamorphosis to the first juvenile. We therefore consider the settling stages of the Achelata (nisto, puerulus) as early juveniles with some initially persisting (mainly behavioral) larval traits rather than as true larvae (compare with the section Presence of larval specific features above). Likewise, the postembryonic stage IV of the Astacidea (sometimes referred to as a zoea or a decapodid) represents an early juvenile rather than a decapodid. This implies that the Achelata and Astacidea lack a decapodid stage. The same may apply to the Euphausiacea, the Amphionidacea, and the decapod groups Dendrobranchiata, Caridea, and Stenopodidea. All these shrimp-​like Eucarida develop through a transitional phase, with gradual morphological and behavioral changes from a late zoea to an early juvenile, passing through several molts. These stages combine typical traits of a late planktonic zoea (well-​developed natatory exopods on the thoracopods) with those of benthic juveniles and adults (swimming with functional pleopods; use of the endopods of the pereopods as walking legs). The exopods of the pereopods are initially natatory, as in the preceding zoeal stages, but then the pleopods gradually take over the swimming function and the pereiopodal exopods are reduced. Nonfunctional vestiges persist sometimes in the early juvenile instars. Concomitantly, the pereopods are increasingly used for benthic walking, as in later juveniles and adults. Hence, these groups show no metamorphosis from a late larval stage to a clearly defined, morphologically and behaviorally different juvenile (i.e., no posterior shift of propulsion, as found in many nonmalacostracan larvae). We therefore discourage using the term decapodid for these taxa to avoid confusion and facilitate comparison between different larval descriptions. Because the number of transitional instars may vary, even among conspecific individuals (Anger 2001), no clearly defined number of larval stages but only an observed range should be given for such taxa. Postlarva The term postlarva is highly ambiguous because it implies a nonlarval nature. In the literature, it has been applied inconsistently to late larval and early juvenile stages (for discussion, see Williamson 1969, Kaestner 1980, Anger 2001). We strongly discourage the use of this term for any larvae.

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PATTERNS OF POSTEMBRYONIC DEVELOPMENT IN CRUSTACEA Anamorphosis, Epimorphosis, and Metamorphosis In Crustacea, the processes of postembryonic development are generally considered to comprise either gradual (anamorphic) or abrupt (metamorphic) changes in morphology, behavior, ecology, and other important biological traits at each ecdysis/​molt (Martin 2014). Strictly speaking, anamorphosis refers to postembryonic patterns of development with a stepwise addition of abdominal body segments at molting, accompanied by otherwise minor morphological changes in body shape or the complexity of appendages (Minelli and Fusco 2013). Many larval as well as all juvenile and adult crustaceans, however, show actually an epimorphic pattern (sensu Enghoff et al. 1993), when molts are accompanied by growth and minor (or no) morphological changes, but not by segment addition (see also Minelli and Fusco 2013). Some authors apply the term epimorphosis to also describe the process of limb regeneration (e.g., Das 2015) or as a term to describe the evolutionary process of a heterochronic shift in larval development toward later, more advanced hatching (e.g., Minelli 2005). For further discussion of terminology, see Chapter 9 in this volume. For convenience, we follow the ACL here and do not distinguish between ana-​and epimorphosis in crustacean larvae (with or without addition of pleomeres, respectively). Both modes are characterized by little morphological change and both may occur within the same developmental phase (e.g., in many Decapoda between successive zoeal stages). Zoeal hatching normally takes place with five pleomeres; the sixth (last) one is added only at the second or third larval molt. All later zoeal molts occur, again, without any addition of body segments. The term metamorphosis denotes a developmental step (molting) with abrupt and conspicuous changes in morphology and other larval traits, normally (but not always) at the transition from the larva to the juvenile/​adult. In Arthropoda, however, this term has been used for seemingly unrelated developmental phenomena of various ontogenetic timing. Also, there may be more than one metamorphosis within a single life cycle. Thompson (1835a,b) described two metamorphoses (double metamorphosis) in the development of Brachyura (zoea–​megalopa, megalopa–​juvenile) and Cirripedia (nauplius–​cypris; cypris–​juvenile) (see, for example, Figs. 6.1A, B; 6.3A; 6.4A; and 6.5A–C). Complete metamorphosis is the most distinctive characteristic of holometabolous insects (all endopterygote orders), which have four developmental phases in their life cycle: embryo, larva, pupa, and adult (imago). Their larval phase is a period of active feeding and growth, whereas the pupal stage is a period of reconstruction. Larval tissues are broken down (histolyzed) and reformed in the adult body plan. The adult phase is a period of dispersal and reproduction. The differences between the types of development in Crustaceans and those in holometabolous insects are remarkable. In holometabolous insects, there is no juvenile phase; the adult stage emerges directly from the pupa (obligate termination of molting) (Enghoff et al. 1993, Hughes et al. 2008). Holometabolous larvae have internalized the primordia of adult structures (imaginal disks). This does not occur in crustaceans, where adult structures are forming gradually, although often through larval stages with a different morphology (e.g., Wolfe and Felgenhauer 1991, Martin 2014, Palero et al. 2014). Internalization of the imaginal disks was probably among the main evolutionary causes and crucial innovations behind the extreme divergence between larvae and adults in holometabolous insects. Another important novelty for holometabolous groups is the pupal stage. In hemimetabolous insects, the last (“adultiform”) nymph stage can be homologized to the pupa (Belles and Santos 2014). In conclusion, the holometabolism concept (implying a complete metamorphosis sensu stricto) cannot be applied to crustaceans without a complete redefinition. The development of crustacean larvae, even in groups with metamorphic molts, is more similar to that of the nymphs in hemimetabolous insects rather than the larvae of holometabolous insects. The fossil record and



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phylogenetic reconstructions suggest that insect holometabolism has evolved only once, with the earliest examples dating back to approximately the mid Carboniferous (Nel et al. 2013). This is obviously independent and much younger than the occurrence of crustacean larvae, which have been found in Cambrian deposits (Fig. 6.2A) (Walossek 1993, Zhang et al. 2010). Hemianamorphosis In most crustaceans, postembryonic development cannot be clearly assigned to either an anamorphic or a metamorphic pattern as proposed in the ACL, but it normally includes both. This combination of different patterns is named hemianamorphosis (Enghoff et al. 1993). It represents the most common mode of larval development in extant Crustacea, followed by purely anamorphic patterns (sensu lato; see ACL; i.e., including epimorphic molts); a minority has a monophasic life cycle without a larval phase (Martin et al. 2014a). Specific examples for fully anamorphic and hemianamorphic patterns of larval development in various crustacean taxa are shown in the following sections. Anamorphic Patterns The anostracans, or fairy shrimp (Branchiopoda), as well as the Cephalocarida show some of the best examples of purely anamorphic developments, without any radical change of morphology at any single molt. This pattern is generally considered ancestral for the Crustacea as fossil larval series from the Lower and Upper Cambrian show a very gradual development (Fig. 6.2A) (Heath 1924, Møller et  al. 2004, Zhang et  al. 2010, Martin et  al. 2014a). Examples of hatching as an orthonauplius and a gradual addition of segments in later stages is found in Artemia spp. and other Anostraca (Fig. 6.2B). The Cephalocarida hatch as metanauplii, showing an equally gradual development. Ostracods also have a very gradual larval development, but start with an unusual bilobed metanauplius (Kesling 1951, Stegner and Richter 2015). Another prime example of anamorphic patterns is found in Caridea, especially in taxa with many (more than five) larval stages (Fig. 6.6). Their earliest zoeal stages are characterized by planktonic behavior, using the anterior thoracopods for swimming; these become mouthparts later in ontogeny. Initially, the posterior thoracic and abdominal appendages (pereopods, pleopods, uropods) are absent. In later zoeal stages, the pereopods develop and increase in size, bearing long natatory exopods, whereas the endopods remain unsegmented, nonfunctional buds that gradually increase in size. Walking behavior begins only in later developmental stages, when the endopods of the pereopods become functional as walking legs, and the pleopods as swimmerets. These semibenthic stages of the Caridea are often referred to as decapodids (i.e., larvae). However, walking with pereopods and swimming with pleopods are also typical traits of juveniles and adults. We therefore consider the postzoeal stages of caridean shrimp as early, morphologically incomplete juveniles with some morphologically visible but functionally insignificant remnants of larval traits (see the “Decapodid” section, this chapter). The size of the endopods of the pereopods increases in successive molts, whereas the exopods are reduced, gradually losing their significance for swimming. Concomitantly, the pleopods gradually take over the natatory function, showing increasing size and setation. In some caridean species, otherwise clearly evident juvenile stages may still exhibit rudimentary exopods on their walking legs as well as later in their development. Hence, the transition from the larval to the juvenile phase of the Caridea is nonmetamorphic and sometimes not evident because adult traits (benthic walking, abdominal swimming) appear already in otherwise larval stages whereas larval traits (natatory exopods) may disappear only gradually during the early juvenile phase (Fig. 6.6). This implies that the “true” (species-​specific) number of larval stages often remains elusive among Caridea.

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Developmental Biology and Larval Ecology Hemianamorphic (“Metamorphic” sensu ACL) Patterns The concept of a metamorphic pattern is commonly used for Crustacea that present one or more metamorphic changes during their postembryonic development (Martin et al. 2014a). Although not widely used, the term hemianamorphic is more correct, as many crustacean larvae also show anamorphosis and/​or epimorphosis. Metamorphic changes occur characteristically between two anamorphic phases—​for example, in the metamorphic molt from the last nauplius of the Cirripedia to a cypris (Fig. 6.1A, B). One of the few known exceptions, with a metamorphic larval molt that is not preceded by an anamorphic phase, is found in some Argulus species (Branchiura). They hatch as a metanauplius, and their first molt is metamorphic, transforming them into a juvenile-​looking animal with a few larval vestiges; all subsequent molts are epimorphic (Møller et al. 2007). The most radical metamorphic molt in the Crustacea is probably the cypris–​kentrogon molt or the ensuing internal molt from the kentrogon to the vermigon (described earlier; Fig. 6.5B, C). The latter entails a loss of basically all external and most internal structures, which transform to a vermiform stage, injected into the host. A typical anamorphic pattern, followed by (in this case, double) metamorphosis, occurs in the cirripedes. During their nauplius phase, the successive stages show only gradual changes (Fig. 6.3B–​G). The subsequent nauplius–​cypris molt is clearly metamorphic (Fig. 6.1A, B). Although the nauplius eye is conserved in the cypris, many other features change dramatically: appearance of a transient complex eye, the single dorsal head shield of the nauplius becomes bilobed, the small and simple first naupliar antennae become highly complex, and four-​segmented organs are used for substratum exploration. In terms of developmental biology, a very interesting change is seen in the transient appearance of natatory thoracopods in the cyprids because these have no anlagen at all in the nauplius (Figs. 6.1B and 6.3B–​G). In all rhizocephalans, this is the only life history stage during which these appendages appear. In all other Cirripedia, they are subsequently lost again when the cypris metamorphoses to a sessile benthic juvenile (Fig. 6.5A–​C), which then grows through numerous epimorphic molts. The euphausiids provide another example of anamorphic patterns with an intercalated metamorphic molt. These shrimp-​like eucarid crustaceans are holopelagic, spending all life history stages in the water column. Their early, fully planktonic larvae are weak swimmers, mostly drifting passively with water currents. The potential for active swimming against currents increases gradually throughout the late larval and the juvenile phases. Consistent with these gradual ontogenetic changes in ecology and behavior, the pattern of morphological developmental is largely anamorphic. The only metamorphic change occurs at the transition from a late nauplius (metanauplius) to the first feeding stage: the calyptopis. The subsequent furcilia stages become increasingly similar to the adults, without showing a metamorphic change of development. Metamorphic transitions from an early to a late larval form, or from the final larval stage to an early juvenile also occur in Achelata, Anomala, Brachyura, and other groups (Harvey et al. 2014, Martin 2014, Palero et al. 2014). At the end of an anamorphic zoeal phase, anomuran or brachyuran crabs molt to a decapodid (Fig. 6.6), which shows dramatic changes in body shape and in the functional morphology of most appendages. The dorsal spine (if present) and the terminal telson spines of the zoea are reduced or lost (Figs. 6.1 and 6.6), the maxillipeds lose their natatory function and become feeding appendages, and the pereopods and pleopods develop into fully functional appendages used for walking and swimming, respectively. These metamorphic changes are particularly striking in Anomala, where the zoeal stages have a slender, shrimp-​like appearance whereas the decapodid is globose, similar to that of the Brachyura. Dramatic changes in morphology and behavior also occur at the molt from the final larval stage to the first juvenile. In Anomala and Brachyura, the transition from a decapodid to a juvenile is less metamorphic than the molt from the penultimate to the final larval stage (zoea–​decapodid) (Anger



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Juvenile / adult phase

Metamorphosis

Larval phase

Embryonic phase

Anamorphosis + Metamorphosis = Hemianamorphosis

Metamorphosis

[2]

[1]

Anamorphosis

Hatching

Morphological development

Epimorphosis sensu stricto

Post-Embryonic Moults Fig. 6.7. Schematic illustration of the principal patterns of postembryonic development in Crustacea. X-​axis: Progressing postembryonic molts marked by short vertical lines (left to right). Y-​axis: Morphological development (continuous scale, bottom to top). (1)  Anamorphosis:  (dark gray) Hatching occurs with a lower than the adult number of body segments. Initially lacking segments are added during the larval phase. Morphological changes, including formation of appendages, occur typically in small steps at each molt until the juvenile phase is reached. This is typical of taxa with a high number of larval stages (\ \ indicates molts not shown). (2) Hemianamorphosis: (light gray) combination of one or more anamorphic phases and one or more metamorphic molts (dramatic morphological changes), followed by epimorphic (sensu stricto) juvenile molts. This is most common pattern in Crustacea. Black dotted vertical lines, transitions between major phases of the life cycle; bold lines, major developmental events. Note that the developmental phase can also be, for example, the zoea phase. See a color version of this figure in the centerfold.

et al. 2015). Despite being morphologically and behaviorally similar to juveniles, however, the presence of natatory pleopods qualifies decapodids as larvae because these appendages disappear subsequently during the molt to a juvenile. In Achelata, the transition from a pelagic phyllosoma larva to a semibenthic puerulus or nisto (here considered as early juvenile stages; described earlier) is also clearly a metamorphic step in ontogeny (Figs. 6.1 and 6.6). Abbreviated Larval Development Many crustaceans pass through a complex life cycle with an abbreviated larval phase. In most cases, this is associated with partial (occurring only in early larval stages) or full lecithotrophy. An abbreviated larval development is characterized by a “shorter duration than that normally seen in the majority of related species in a taxon, and which results in fewer morphologically discrete instars and/​or reduced ontogenetic duration” (Rabalais and Gore 1985, p. 71). Such exceptions from the

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Developmental Biology and Larval Ecology taxon-​specific patterns are most frequently found in species that live and reproduce in physiologically, nutritionally, or otherwise demanding habitats including fresh water, anchialine caves, the deep sea, or polar regions, where planktonic food limitation or stressful physical conditions select against an extended, planktivorous larval phase (Rabalais and Gore 1985, Vogt 2013, Anger 2016). This phenomenon is most common in taxa that normally have an extended larval development with many stages—​for instance, caridean shrimp (e.g., Guerao 1993, Magalhães 2000, Rodriguez and Cuesta 2011). When larval development comprises both an anamorphic phase as well as at least one metamorphic transition (e.g., anomuran and brachyuran decapods; Cirripedia), it is usually only the anamorphic part (in these examples, the zoeal or nauplius phase) that is shortened; the metamorphic part is retained [here, the double metamorphosis zoea–​decapodid–​juvenile; see also (Clark 2005)]. In some exceptional cases among Brachyura, the entire zoeal phase is eliminated (or actually passed within the egg case), so that hatching occurs as a megalopa (Wear 1967), suppressing the anamorphic changes of free-​living larvae but maintaining the metamorphosis of the megalopa to the juvenile. On the other hand, an abbreviated and lecithotrophic zoeal phase can also be directly followed by an early, morphologically incomplete juvenile with vestigial traits of a megalopa, keeping only the anamorphic (or epimorphic sensu stricto) pattern (Gonzalez-​Gordillo et al. 2010). Similar patterns of abbreviated development with very few lecithotrophic zoeal stages followed by an early, morphologically incomplete juvenile (mostly still referred to as a zoea) occur also in caridean shrimp, especially in hololimnetic palaemonids (see the previous section “The Fundamental Question”) (Magalhães 1988, Jalihal et  al. 1993). In Cirripedia, there are examples of species in which the nauplius phase is skipped completely and a cypris larva hatches (e.g., in all freshwater rhizocephalan species as well as all akentrogonids) (Høeg and Møller 2006). An abbreviation of the larval phase thus leads to a heterochronic shift from anamorphic toward increasingly metamorphic patterns (Minelli 2005). The endpoint of this evolutionary trend is direct development—​when an embryo “molts” directly to a juvenile. Intraspecific Variability in Developmental Pathways Patterns of postembryonic development, including the number and morphology of larval stages, are commonly considered as taxon-​specific, fixed characteristics. Although this may be true for the majority of crustaceans with complex life cycles, there are numerous exceptions. Intraspecific variability has been documented for numerous clades, especially groups with extended, fully, or predominantly anamorphic patterns of larval development such as the Euphausiacea (Feinberg et  al. 2006, Antezana and Melo 2008)  and Caridea (Criales and Anger 1986, Wehrtmann and Albornoz 1998), but also Thalassinidea (Strasser and Felder 1999), Anomura (Wang et al. 2007), and Brachyura (Zeng et al. 2004, Guerao et al. 2012). Many authors, especially field workers who are used to studying material collected from plankton samples, tend to ignore intraspecific variability, considering it unimportant or a laboratory artifact caused by suboptimal rearing conditions. However, there is ample evidence that developmental variability is mainly the result of phenotypic plasticity, which allows for flexible responses to variations in nutritional or physical factors (Criales and Anger 1986, Weiss et al. 2010; see also Chapter  10 in this volume). Such variations in environmental conditions occur in the natural plankton as well (normally even more than in controlled laboratory cultures), where they most probably exert similar effects. This is also supported by observations of variable developmental pathways in field studies (Strasser and Felder 1999, Wehrtmann and Albornoz 2003). Moreover, the patterns of development can vary among different broods, reflecting genetic factors and/​or carryover effects of environmentally influenced female conditions (Giménez et al. 2004), and even among sibling larvae reared under identical conditions (individual variability, probably based on genetic variation) (Anger 2001).



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Intraspecific variability in the number and duration of larval stages is associated with variations in larval morphology, body size, biomass, and physiological condition in a given instar (i.e., after a determined number of molts). On the other hand, these traits can be similar in different instars and after differential periods of development. All this means is that the rates of molting and those of morphological and physiological development are not strictly correlated in taxa with intraspecific variability. Consequently, predictions of larval morphology, size, and so on, from the number of molts (precisely known only in laboratory studies) are not always possible. Likewise, a particular morphological stage found in plankton samples does not always allow for a precise inference on the number of previous molts or the time passed since hatching. All this has implications for modeling studies describing and predicting patterns of larval dispersal, recruitment, and genetic exchange between populations (e.g., Pires et al. 2013, Anger et al. 2015). Such models are based on the occurrence of particular morphological stages of larvae in plankton samples, in conjunction with hydrographic data (water temperatures, stratification, direction and speed of currents), known relationships between physical factors and rates of molting, as well as knowledge of larval behavior (e.g., vertical migrations). Although such models should be fairly reliable among taxa with little or no variability in developmental patterns, researchers must be aware of biological variability as a potential source of uncertainty. Developmental Patterns and Phylogenetic Significance In most studies, the subphylum Crustacea is paraphyletic and the hexapods are derived from crustacean ancestors. A monophyletic taxon Crustacea + Hexapoda, Tetraconata [Dohle (2001); originally Pancrustacea (Zrzavy et al (1997) and Štys 1997)], is well supported by phylogenies that are based on molecular data, morphological characteristics, and large phylogenomic analyses (e.g., Giribet et al. 2001, Oakley et al. 2013). However, the inference of the homologies between hexapods and crustaceans is difficult because the basal divergence within Tetraconata occurred no later than 530 million years ago (Regier et al. 2005). In the great majority of the hexapods (all insects), postembryonic development is normally epimorphic. However, uniquely among hexapods, the Protura show hemianamorphic development whereby somites are added during molts (Pass and Szucsich 2011). Given that some evidence indicates that the clade Entognatha (Protura + Diplura + Collembola) is a sister group of all other hexapods (class Insecta), probably the anamorphosis of proturans is a plesiomorphic character shared by crustaceans. In addition, like the crustaceans, the Entognatha (Hexapoda) and wingless insects (Archaeognatha and Thysanura) continue to molt and grow as adults while new appendages develop on the abdomen. However, it still remains unclear which of the major crustacean groups represents the sister group of Hexapoda (e.g., see Rota-​Stabelli et al. 2011, von Reumont et al. 2012). Oakley et al. (2013) proposed a new clade, Allotriocarida (which includes Hexapoda, Branchiopoda, Remipedia, and Cephalocarida), which has also found support in newer studies. Currently, the best candidate for a sister group to the Hexapoda seems to be the Remipedia, which was highly supported in a recent study by Schwentner et al. (2017) using the first transcriptomics data and multimodel testing.

CONCLUSIONS True to the highly variable biology and morphology of the Crustacea, the patterns of free-​living larval development are as equally both variable and complex. Being an important link between life cycle phases, larval diversity has received a lot of attention from many researchers. This chapter provides an overview of the most important patterns found in free-​living, prejuvenile life history stages (i.e., larvae) based on examples from various groups.

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Developmental Biology and Larval Ecology In the literature dealing with crustacean larvae, many terms such as stage, instar, or phase are used ambiguously, following different and often poorly explained definitions. This terminological uncertainty includes even the basic term larva itself. We suggest here that a mere lack, or underdeveloped condition, of juvenile–​adult traits (negative criterion) is insufficient to consider a postembryonic life history stage as a larva. Instead, we propose that a larva must show specific traits that are lacking in the other (embryonic, juvenile, adult) phases of the life cycle (positive criterion). In at least some species in which a strongly abbreviated larval development has been reported, we suggest a direct development (i.e., a complete lack of a larval phase). Reviewing the names of specific larval stages in the Decapoda (e.g., zoea, decapodid), we also show that these names are used inconsistently. This terminological variability in the literature reflects the diversity of larval forms both between and within crustacean groups, as well as diverging views among research schools. For the sake of comparability, and to reduce or (hopefully) eliminate such problems of inconsistency (and often redundancy) in the use of larval names, we suggest that authors always provide (and justify) their own terminology, or at least refer to previously published work where such definitions are given. Besides variability in terminology, we draw your attention to the intraspecific variability in developmental pathways. This variability is caused by the variable degrees of morphological change after each molting cycle (including lack of such changes, termed mark-​time molting). This confounding phenomenon has frequently been ignored or misinterpreted as laboratory artifacts, although it is quite common in some crustacean groups (e.g., euphausiids, caridean shrimp). We therefore suggest that its occurrence should always be described as a species-​specific, developmental trait. This implies that many species do not have a clearly defined number of distinct larval stages and that larval morphology is not strictly linked to a particular number of postembryonic molts. Likewise, the morphological and behavioral transition from the larval to the juvenile phase occurs in some groups (e.g., Caridea) as a gradual (anamorphic) process, often through a variable, high number of molting cycles with only small changes at each molt. Both larval and early juvenile traits may coexist transitorily, which implies that, in such cases, no species-​specific number of larval stages can be determined. Proposing a robust and simple two-​mode system (hemianamorphic and anamorphic), presented in Fig. 6.7, we attempt to outline our idea of the most general developmental patterns in all Crustacea with free-​living larvae. Almost all known free-​living larval patterns can be categorized by this system, including very special cases such as mark-​time molting in some Decapoda and the double metamorphosis in the Rhizocephala. Because it focuses on one of the most important drivers of evolution in the Crustacea, the ubiquitous molting cycle, it avoids the complexity of the inherent heterochrony (relative developmental timing) of the different developmental pathways in different species while still allowing for the intraspecific variability mentioned earlier. In summary, we caution against very strict and comprehensive terminological systems that attempt to describe virtually all larval forms and developmental patterns in the Crustacea. Rather, we encourage authors to show flexibility for exceptions (e.g., describing the gradual transition from the larval into the juvenile phase in some taxa), rather than always providing yet another fixed species-​ specific number of larval stages.

REFERENCES Akther, H., M.D. Agersted, and J. Olesen. 2015. Naupliar and metanaupliar development of Thysanoessa raschii (Malacostraca, Euphausiacea) from Godthåbsfjord, Greenland, with a reinstatement of the ancestral status of the free-​living Nauplius in Malacostracan evolution. PLoS One 10:e0141955.



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Developmental Biology and Larval Ecology Tully, O., and D.T. Nolan. 2002. A review of the population biology and host–​parasite interactions of the sea louse Lepeophtheirus salmonis (Copepoda: Caligidae). Parasitology 124:S165–​S182. Vogt, G. 2013. Abbreviation of larval development and extension of brood care as key features of the evolution of freshwater Decapoda. Biological Reviews 88:81–​116. Vogt, L., P. Grobe, B. Quast, and T. Bartolomaeus. 2012. Accommodating ontologies to biological reality: top-​ level categories of cumulative–​constitutively organized material entities. PLoS One 7:e30004. 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. Walker, G. 2001. Introduction to the Rhizocephala (Crustacea: Cirripedia). Journal of Morphology 249:1–​8. Walossek, D. 1993. The upper Cambrian Rehbachiella and the phylogeny of Branchiopoda and Crustacea. Fossils and Strata 32:1–​202. Walsh, C.J. 1993. Larval development of Paratya australiensis Kemp, 1917 (Decapoda: Caridea: Atyidae), reared in the laboratory, with comparisons of fecundity and egg and larval size between estuarine and riverine environments. Journal of Crustacean Biology 13:456–​480. Wang, F.L., H.L. Hsieh, and C.P. Chen. 2007. Larval growth of the coconut crab Birgus latro with a discussion on the development mode of terrestrial hermit crabs. Journal of Crustacean Biology 27:616–​625. Wear, R.G. 1967. Life-​history studies on New Zealand Brachyura: 1. Embryonic and post-​embryonic development of Pilumnus novaezealandiae Filhol, 1886, and of P. lumpinus Bennnett, 1964 (Xanthidae, Pilumninae). New Zealand Journal of Marine & Freshwater Research 1:482–​535. Wehrtmann, I.S., and L. Albornoz. 1998. Larval development of Nauticaris magellanica (A. Milne Edwards, 1891) (Decapoda: Caridea: Hippolytidae), reared under laboratory conditions. Bulletin of Marine Science 62:45–​72. Wehrtmann, I.S., and L. Albornoz. 2003. Larvae of Nauticaris magellanica (Decapoda: Caridea: Hippolytidae) reared in the laboratory differ morphologically from those in nature. Journal of the Marine Biological Association of the United Kingdom 83:949–​957. Weiss, M., S. Thatje, and O. Heilmayer. 2010. Temperature effects on zoeal morphometric traits and intraspecific variability in the hairy crab Cancer setosus across latitude. Helgoland Marine Research 64:125–​133. Williams, J.D., and C.B. Boyko. 2012. The global diversity of parasitic isopods associated with crustacean hosts (Isopoda: Bopyroidea and Cryptoniscoidea). PLoS One 7:e35350. Williamson, D.I. 1969. Names of larvae in the Decapoda and Euphausiacea. Crustaceana 16:210–​213. Williamson, D.I., and D.E. Bliss. 1982. Larval morphology and diversity. Pages 43–​110 in L.G. Abele, editor. Embryology, Morphology and Genetics. Academic Press, New York, New York. Wilson, G.D.F., C.A. Sims, and A.S. Grutter. 2011. Toward a taxonomy of the Gnathiidae (Isopoda) using juveniles: the external anatomy of Gnathia aureamaculosa Zuphea stages using scanning electron microscopy. Journal of Crustacean Biology 31:509–​522. Wolfe, S.H., and B.E. Felgenhauer. 1991. Mouthpart and foregut ontogeny in larval, postlarval, and juvenile spiny lobster, Panulirus argus Latreille (Decapoda, Palinuridae). Zoologica Scripta 20:57–​75. Zeng, C.S., S.J. Li, and H. Zeng. 2004. Occurrence of additional zoea-​VI larvae in the mud crab, Scylla paramamosain (Estampador), reared in the laboratory. Hydrobiologia 529:49–​58. Zhang, X., A. Maas, J.T. Haug, D.J. Siveter, and D. Waloszek. 2010. A eucrustacean metanauplius from the Lower Cambrian. Current Biology 20:1075–​1079. Zrzavý, J., V. Hypša, and M. Vlaskova. 1997. Arthropod phylogeny: taxonomic congruence, total evidence and conditional combination approaches to morphological and molecular data sets. Pages 97–107 in R. A. Fortey and R. H. Thomas, editors. Arthropod Relationships. Chapman and Hall, London, UK.

7 EFFECTS OF ENVIRONMENTAL CONDITIONS ON LARVAL GROWTH AND DEVELOPMENT

Chaoshu Zeng, Guiomar Rotllant, Luis Giménez, and Nicholas Romano

Abstract The vast majority of crustaceans are aquatic, living in either marine or freshwater environments. Marine crustaceans—​such as copepods, in particular—​are ubiquitous in the oceans and perhaps the most numerous metazoans on Earth. Because crustaceans occur in all marine habitats, their larvae are exposed to highly diverse and sometimes variable environmental conditions, including extreme situations in which various environmental factors exert significant effects on larval growth and development. This chapter first describes the effects of food availability on crustacean larvae. Food paucity is a commonly occurring scenario in the wild, which can directly affect larval growth and development and, in severe cases, results in mortality. In the subsequent sections, we cover the effects of temperature and salinity—​the two most prominent physical parameters in the aquatic environments—​on growth and development of crustacean larvae. We then discuss the influence of other important physicochemical factors in aquatic environments on larval growth and development, including dissolved oxygen, light, ocean acidification, and pollutants. Finally, the last two sections of this chapter discuss synergistic effects of different environmental factors and suggest future research directions in this field.

INTRODUCTION Crustaceans inhabit a wide range of ecosystems globally; hence, they are exposed to highly diverse environmental conditions, including extreme conditions, such as those at high latitudes and in the

Developmental Biology and Larval Ecology. Edited by Klaus Anger, Steffen Harzsch, and Martin Thiel. © 2020 Oxford University Press. Published 2020 by Oxford University Press.

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Developmental Biology and Larval Ecology deep sea. This chapter describes the effects of various environmental factors, including food availability, temperature, salinity, dissolved oxygen, light, and pollutants, on the growth and development of crustacean larvae. To cope with the highly diverse conditions they are likely to encounter, crustaceans with complex life cycles have evolved into a wide variety of life history adaptations. For instance, the number of larval stages and their duration differ vastly not only among taxonomic groups, but also within a taxonomic group (Olesen 2018). The variations range from direct or abbreviated development to very long larval durations of more than nine months with more than 20 molts (e.g., rock lobsters; for details, see Chapter 6 in this volume). Larval growth is most commonly quantified as a change in body size or weight over a defined period of time, which is usually the duration of a specific developmental stage. On the other hand, development is defined as the time required for a newly hatched or newly molted larva to reach a designated subsequent developmental stage during larval ontogeny (Anger 2001). Suboptimal environmental conditions generally reduce larval growth and affect their development first; however, if the conditions are severe and/​ or become prolonged, it could threaten larval survival, ultimately leading to mortality. Hence, although larval growth and development are the main themes discussed in this chapter, the effects of environmental conditions on larval survival are also discussed when necessary. Crustacean larval development generally encompasses discrete stages punctuated by molting events. Various environmental factors can all affect the larval molting cycle, but some (e.g., temperature) exert a greater influence on molting frequency or larval development whereas others (e.g., food availability) appear to affect more significantly larval size increments during ecdysis or larval growth (Anger 2001). To evaluate the effects of environmental parameters on larval growth, various indices have been used. Of these indices, the most common ones are based on body size and biomass; however, other indices have been also increasingly used; they include elemental and proximal compositions, lipid classes and their contents, digestive and metabolic enzyme activity, as well as DNA and RNA and their ratio (e.g., Ritar et al. 2003, Rotllant et al. 2010, Guerao et al. 2012, Wu et al. 2014). Most studies on development and growth of crustacean larvae carried out in the laboratory focus on larval responses under constant conditions (Anger 2001). Data from field studies, however, show that growth rates in natural environments may differ from those in the laboratory under assumedly similar conditions (e.g., González-​Gordillo and Rodríguez 2000, Guerao et  al. 2008, Westphal et al. 2014). Such discrepancies are not surprising because, in laboratory experiments, other major environmental variables are usually maintained unnaturally constant to evaluate the effects of one or a few factors under investigation. Although laboratory experiments inevitably create unnatural conditions, field studies—​on the other hand—​are limited by an incomplete knowledge of both temporal and spatial changes of prevailing environmental conditions. Moreover, it is difficult to pinpoint the influences of a particular factor under investigation when other environmental factors are undergoing simultaneous fluctuations, as is the case with field studies. Hence, a more holistic approach to understanding the effects of environmental factors on larval growth and development requires the combination of both laboratory and field studies, thus pulling together their complementary strengths. The main environmental factors modifying the growth and development of crustacean larvae include food availability and quality (the latter aspect is discussed in detail in Chapter 11 in this volume), temperature, and salinity. Furthermore, other physicochemical factors such as dissolved oxygen, pH, light, and pollutants can also exert major effects. As these environmental factors are often covarying, their combined effects have received increasing attention more recently, specifically in relation to research on climate change (e.g., Walther et al. 2010, Whiteley 2011, Agnalt et al. 2013, Gonzalez-​Ortegón et al. 2013, Wood et al. 2015). One section of this chapter is hence devoted to discuss this important new trend. Finally, the direction of future research is recommended in the last section of this chapter.



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FOOD AVAILABILITY For the vast majority of crustacean larvae, successful development is only initially dependent on yolk. When yolk becomes depleted, the intake of food becomes crucial for the remaining time of larval development (Anger 2001, Wouters et al. 2001). The degree of dependence on yolk reserves for successful larval development differs substantially among crustacean species. For example, larvae of portunid (e.g., Andrés et al. 2010) and most majid crabs (e.g., Anger and Dawirs 1981, Guerao et al. 2012) are planktotrophic; they consume food immediately upon hatching, and starvation prevents them from molting to the next larval stage. On the other hand, newly hatched larvae of some crustaceans are obligatory lecithotrophs. In other words, they do not immediately feed after hatching; instead, they rely solely on yolk reserves for nutrition as they develop through the initial instars (Urzúa et al. 2013). Examples of obligatory lecithotrophy include nauplii of various marine shrimp and copepods. In addition, there are crustacean larvae that exhibit facultative lecithotrophy. These larvae are also capable of successful development to the next instar in the complete absence of food, but feed when food is available. Facultative lecithotrophy is found not only in newly hatched larvae, such as the first zoeae of marine cleaner shrimp belonging to the genera Stenopus and Lysmata (e.g., Figueiredo and Narciso 2006), but also in later larval stages, such as the second-​stage zoeae of the freshwater prawn Macrobrachium amazonicum (Anger and Hayd 2009) and the megalopae of some freshwater crabs from the genera Sesarma and Metopaulias (Anger and Schubart 2005). In facultative lecithotrophic larvae, larval biomass generally increases or decreases, depending on whether food is consumed. When food is absent, survival is normally less, but development is faster compared to fed larvae (Anger and Schubart 2005). In nature, crustacean larvae are capable of consuming a broad range of food items, from microplankton, phytoplankton, to various zooplankton (Harms et al. 1994, Shaber and Sulkin 2007); and some larvae may also consume nano-​and picoplankton (Thompson et al. 1999, Finlay and Roff 2004). Larval size often determines their trophic position in planktonic food webs, with many species changing their trophic level during ontogenetic development, which was confirmed by stable isotope studies (Le Vay et al. 2001, Le Vay and Gamboa-​Delgado 2011). During times of food scarcity, some crustacean larvae may cannibalize conspecifics, which serves as an important strategy known as a lifeboat mechanism (van den Bosch et al. 1988). For instance, the grapsid crab Armases miersii releases its larvae at different times in isolated supratidal pools characterized by unpredictable food availability. In such pools, the coexistence of different larval stages allows larvae at advanced stages to cannibalize younger conspecifics when food is scarce. Such a mechanism ensures some larvae could successfully develop and become juveniles (Anger 1995). As a result of the often patchy distribution of plankton found in nature, along with diel vertical migrations undertaken by many crustacean larvae, the quantity and quality of natural food for developing larvae are highly variable and unpredictable; periodic starvation is likely to occur. If starvation is prolonged, the point of no return (PNR) may be reached. The PNR is defined as the point in time when the capacity of larvae to recover from nutritional stress is lost as a result of irreversible damages caused by starvation. Consequently, the larvae eventually die without molting to the next instar, even when food becomes available again and is ingested by the larvae (Anger and Dawirs 1981, Anger 2001). Figure 7.1 shows a conceptual model of PRN: larval resistance to initial starvation is quantified through PNR50 (i.e., the initial starvation time leads to 50% of larvae losing their ability to develop to the next instar, despite the resumption of feeding). PNR50 is species specific. For example, PNR50 is reportedly 3.8 days (Zhang et al. 2015) and 4.6 days (Liddy et al. 2003) at 25°C for newly hatched larvae of the prawn Exopalaemon carinicauda and the western rock lobster Panulirus cygnus, respectively; whereas for the spider crab Maja brachydactyla, it is 2.8 days at 18° C (Guerao et al. 2012). Temperature can significantly influence PNR50; in newly hatched larvae of the shore

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crab Carcinus maenas, PNR50 is almost halved at 18°C compared to 12°C, likely a result of increased catabolism under the higher temperature of 18°C (Dawirs 1984). Adaptations to patchy food environments allow crustacean larvae to complete larval development despite limited access to food. Among larvae performing diel vertical migration, it is expected that access to abundant prey is limited to a few hours during the nighttime, when larvae are in presumably food-​rich surface layers. Laboratory experiments have shown that decapod larvae can develop to the next instar and sometimes complete their full larval phase under a limited access to prey of only four to six hours per day (Sulkin et al. 1998, Gimenéz and Anger 2005, D’Urban Jackson et al. 2014, González-​Ortegón and Giménez 2014). Such an ability was seen in larvae of the European lobster Homarus gammarus and the common prawn Palaemon serratus over a wide range of temperatures (González-​Ortegón and Giménez 2014, D’Urban Jackson et al. 2014). Under starving conditions, the main energy stores (i.e., proteins, carbohydrates, and lipids) are catabolized by the larvae for energy to sustain life (Anger 2001, Ritar et al. 2003). Anger (2001) described a general pattern of energy use in starved decapod larvae: During short-​term food deprivation, accessible energy-​rich lipid reserves are preferentially mobilized, but when much of the accessible lipid pool has been depleted, proteins are increasingly used, as reflected in the degradation of structures such as muscle and nervous tissue. It is also known that a significant part of the lipid pool is bound in crucial cell structures such as cell membranes, which normally is unavailable for energy metabolism although during the final phase of starvation before death, such structural lipids may also be degraded and become available for energy. This metabolic shift has been considered



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as the underlying cause of the PNR, because excessive catabolism of structure lipids likely induces irreversible damages (Anger 2001, Abrunhosa and Kittaka 1997). However, more recent studies indicate that energy use in starved decapod larvae may not always fit into this general pattern. For instance, protease activity was found to be greater in starved larvae of the spiny lobster Jasus edwardsii ( Johnston et al. 2004), the spider crab Maja brachydactyla (Rotllant et al. 2010), and the mud crab Scylla serrata (Genodepa 2015) compared to the fed larvae, which instead had greater lipase or esterase activities. In fact, greater amylase activity was also detected in the starved larvae of S. serrata when compared to the fed ones (Genodepa 2015). On the other hand, a general reduction in digestive enzyme activities as a result of food deprivation was reported in C. maenas larvae (Harms et al. 1994). The pattern of nutritional reserve use under starvation may also depend on the level of larval foraging activity. For instance, in the newly hatched larvae of the blue swimmer crab Portunus pelagicus, the initial high level of foraging activity observed was suggested as the likely driver of the highest percentage loss of dry weight (17%) and total lipids (38%) during the first day of a three-​ day starvation period (average duration of zoea I larvae when fed, approximately three days). Such activity was subdued on subsequent days, likely as a strategy to reduce energy expenditure as nutrition reserves rapidly deplete (Wu et al. 2017). It was also found that phospholipids dominated the lipid constituents in the newly hatched zoea I larvae of P. pelagicus, but decreased by approximately 50% by the end of the three-​day starvation, suggesting that a substantial portion of these membrane structural lipids had been oxidized for energy during extended starvation, which likely led to the PNR (Wu et al. 2017).

TEMPERATURE Most experimental evidence suggests that the growth of crustacean larvae is at a maximum near the temperature at which the physiological performance of a given larval stage is optimal and growth decreases at both higher and lower temperatures. On the lower end of the tolerance range, it seems that a reduction in the instantaneous rate of biomass accumulation is stronger than the deceleration of development whereas at unfavorably high temperatures, the enhancement of biomass growth appears to be relatively weaker than the concurrent acceleration of molting frequency (Pörtner 2001). Pörtner (2001) proposed that thermal tolerance windows should be defined as lower and upper pejus temperatures (Tp) and critical temperatures (Tc)—​in other words, the upper and lower Tp describe the range at which aerobic scope is maximal and the functioning of the organism is unrestrained, which are estimated based on direct observations of animal activity. The transition from aerobic to anaerobic metabolism is defined by Tc limits that are estimated based on the relationship between standard metabolic rate and temperature. Anaerobic metabolism provides substantially less energy than aerobic metabolism; hence, less energy is likely to be invested for growth (Pörtner 2001). In a study by Storch et al. (2011) on larvae of the Chilean kelp crab Taliepus dentatus, ontogenetic changes in Tc limits were identified with a narrower range for megalopae compared to the zoeal larvae. This was consistent with the hypothesis that thermal tolerance should be lowered as organismal complexity increases. The narrowing of thermal tolerance limits, as larvae developed from zoea I to the megalopal stage, was consistent with the patterns of larval oxygen consumption, suggesting that zoea I  larvae might be more tolerant to lower dissolved oxygen than megalopae (Fig. 7.2). It was also observed that thermal tolerance varied across early life history stages in the porcelain crab Petrolisthes cinctipes. That is, larval stages that were expected to experience more variable temperatures showed a higher thermal tolerance than those inhabiting less thermally variable habitats (Miller et al. 2014). Such a pattern, however, was not observed in a closely related

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Fig. 7.2. Temperature limits in megalopae of the Chilean kelp crab Taliepus dentatus. (A). The pejus temperature (Tp) was established using pleopods beats. The decrease in beats indicate a clear decrease in performance. (B) The critical temperature (Tc) was established based on changes in reductions of oxygen consumption rates, which is quantified as the difference between the active metabolic rate (AMR) and the standard metabolic rate (SMR). The AMR was measured directly; the SMR was calculated from the difference between AMR and costs of pleopod beating. Such costs were estimated from regressions between pleopod beatings and AMR. Note the decreased pleopods beats and oxygen consumption rates at Tc. Modified from Storch et al. (2011), with permission from © Inter-​Research 2011.

species, Petrolisthes manimaculis, living in colder habitats (Miller et al. 2014). Similarly, an ontogenetic shift in stage-​specific temperature optimum over successive larval stages, as determined by maximum biomass achieved, has been reported for various crustacean larvae. For instance, the dry biomass of spiny lobster Sagmariasus verreauxi phyllosoma was reportedly significantly greater when cultured at 25°C, whereas for pueruli, a greater biomass was attained at 21°C (Fitzgibbon and Battaglene 2012). Within a thermal tolerance range, the duration of larval development for a given stage generally decreases with increasing temperature, which is independent of the number of larval stages [Giménez (2011) and references therein]. For example, when the effect of temperature on larval development of the crab M.  brachydactyla was examined between 15°C and 30°C, a complete larval development was only observed up to 24°C. The shortest duration of 17 days was recorded at 21°C, whereas it took substantially longer (27 days) at 15°C (Castejón et al. 2015). Similarly, for the lobster H. gammarus, larval duration decreased by half, from 26 days to 13 days as temperature increased from 14°C to 22°C (Schmalenbach and Franke 2010). Likewise, an increase in temperature from 10°C to 15°C decreased the postembryonic development time by nine days in the copepod Eurytemora affinis (Devreker et al. 2007). Temperature has also been reported to affect larval instar number. The shrimp Palaemon macrodactylus showed an increase in larval instar numbers from five to seven at lower temperatures



Larval Growth and Development

(Vázquez et  al. 2015). Temperature difference may also explain differences in larval instar numbers found between various populations of the same species. For instance, larvae of the shrimp Crangon crangon from the warmer Baltic Sea usually developed to the juvenile stage through four or five instars, whereas those from the colder North Sea generally passed through five or six instars (Criales and Anger 1986). In species with a wide latitudinal cline and thermal tolerance, larval development rates between different populations may also differ at similar temperatures, and larger larvae have frequently been observed in colder parts of their biogeographic range. For example, larvae of the coastal shrimp Palaemon serratus from the warmer Mediterranean had a faster development rate but smaller size than those from Ireland (Kelly et al. 2012). At 9°C, the development of spider crab Hyas araneus megalopae from the Arctic Svalbard population took 10 days longer than those of the North Sea population (40 days vs. 30 days), whereas dry weight of the former was significantly higher (Walther et al. 2010). Similarly, the copepodites of Temora longicornis and Acartia clausi from populations acclimated to cold temperatures attained higher weights than those from the warm-​acclimated populations (Leandro et al. 2006, Dzierzbicka-​Glowacka et al. 2011). The relationship between size and temperature follows the temperature–​size rule: Within an optimum range, larger body weights of newly settled juveniles generally occur at lower temperatures. This has been reported in various crustaceans, including both decapods (e.g., Dawirs et al. 1986, Künisch and Anger 1984, Jackson et  al. 2014)  and copepods (Leandro et  al. 2006, Forster et  al. 2011b). However, in brine shrimp Artemia early larvae, such a trend was not found (Forster and Hirst 2012). In fact, in a meta-​analysis, Forster et al. (2011a) found that thermal responses in eight crustacean species were weaker during early larval stages when compared to advanced stages, pointing to important ontogenetic effects. The length and width of different body parts of crustacean larvae can also be affected by temperature. A larval morphometric approach has been used to identify the optimal temperature (20°C) for the king prawn Melicertus latisulcatus; the larvae of the species reared at temperatures from 17°C to 25°C showed differences not only in body length and width, but also in the length of the antenna, carapace, and abdomen (Rodgers et al. 2013). In a similar way, at low temperatures, prosome lengths of A. clausi copepodites were shorter in the northern population (warmer waters) when compared to those of the southern population (Leandro et al. 2006). Weiss et al. (2010) also reported that early zoeae of the hairy crab Romaleon setosum from a southern colder region [sea surface temperature (SST), 10–​16°C] had a significantly longer total length, including those of the rostral and dorsal spine, than those from a northern warmer region (SST, 16–​20°C) or of the larvae reared at 24°C. In addition, deformations were found in those larvae reared at 24°C, suggesting that 24°C—​a temperature coinciding with strong El Niño events—​is near the upper thermal tolerance limit of the larvae. In crustacean larvae, temperature acclimation responses, which involve the process of adjusting to temperature changes in relatively short time periods within the lifetime of the organism (as opposed to adaptation that takes place over generations) are not always beneficial. In fact, this may cause a malfunction in environments where temperature changes rapidly or when seasonal changes are unpredictable, such as during El Niño events. Moreover, thermal acclimation ability can vary between populations from different geographic regions for a species, which was observed in larvae of the gooseneck barnacle Pollicipes elegans. Larval P. elegans from the northern hemisphere (Mexico) that already adapted to recurrent seasonal temperature changes were more resistant to temperature increases than those from the Peruvian coast in the southern hemisphere (Crickenberger et al. 2015). In conclusion, although differences in temperature could affect both growth and development of crustacean larvae, temperature is thought to exert more prominent effects on larval development.

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SALINITY Salinity is a measure of the quantity of dissolved salts in the water and it can vary from around 35 in the open oceans to less than 0.5 in freshwater systems. For some crustaceans, particularly decapods, the life cycle involves spending a part or the whole larval phase within estuaries where salinity often fluctuates widely. In other cases, larvae undergo long-​distance dispersal/​recruitment migrations during which salinity changes are also commonly encountered (Bermudes and Ritar 2005, Torres et al. 2011, Criales et al. 2015). Because larvae are generally weak swimmers, their ability to avoid osmotically stressful conditions can be limited, which could lead to changes in their physiology, energetic reserves, behavior, and distribution (Anger 2003, Vilas et al. 2006). Because osmotic stress affects larval survival, growth, and development negatively, various strategies have been developed to cope with such challenges, which include vertical migrations (i.e., horizontal larval exports using tidal currents), diadromous migrations of reproductive-​ready females, and osmoregulation (Torres et al. 2011). Such different strategies may be adopted under different situations by different species or life stages. For instance, when a rapid and short-​term decrease in salinity occurs, larvae can reduce their swimming activity and thus sink deeper to avoid low-​salinity surface waters until conditions change (Queiroga and Blanton 2004). In the case when seasonal rainfall drastically reduces salinity in estuaries, migration to the open ocean where salinity is more stable is another strategy adopted by some larvae (Diele and Smith 2006, Torres et al. 2011), particularly osmoconformers (i.e., larvae with internal osmolality similar to that of the external environment resulting from a lack of osmoregulatory ability). Most decapod larvae are osmoconformers; their small size and relatively thin cuticle limit their ability to maintain a stable internal osmolality at varying salinities. It is important to stress, however, that despite their similar osmolality to the environment, the internal ion composition of osmoconformer larvae may differ substantially from that of the environment (Burton 1992). This is often accomplished by the adjustment of their free amino acid content through protein catabolism/​synthesis, which acts as organic osmolytes, to higher levels at high salinities and lower levels at low salinities to reduce the passive influx/​efflux of ions across cell membranes. Such a mechanism is known as intracellular isosmotic ion regulation (Charmantier et al. 2009) and has been demonstrated in megalopae of the blue crab Callinectes sapidus (Burton 1992), nauplii of the copepod Apocyclops panamensis (Lindley et al. 2011), and during larval ontogeny of H. gammarus (Haond et al. 1999). On the other hand, extracellular osmoregulation involves the active movements of ions into or out of the hemolymph, depending on external salinity, which is often fueled by Na+/​K+-​adenosine triphosphatase (ATPase) activity. The stronger the capability, the more independent the internal osmolality can be maintained regardless of the environmental salinity changes. Although extracellular osmoregulation is generally more common or stronger in juvenile and adult crustaceans (Romano and Zeng 2012), some crustacean larvae have been demonstrated to have such an ability. For example, all larval stages of the neotropical crab Armases miersii have been shown to express strong hypo-​osmoregulation capacity at high salinities as well as hyperosmoregulation at low salinities (Charmantier et al. 1998). Although osmoregulation is costly energetically, a greater osmoregulatory ability allows for a retention strategy and ensures sustained larval growth and development under low salinities (Torres et al. 2011). The osmoregulatory abilities of crustacean larvae may change with ontogenetic development and this is often related to the pattern of their ontogenetic migration (Charmantier et  al. 2002, Cieluch et al. 2004, Torres et al. 2011, Lignot and Charmantier 2015). For example, Charmantier et  al. (2002) reported that the first-​stage zoeae of the South American saltmarsh crab Neohelice granulata were able to hyperosmoregulate, which coincides with the fact that these larvae were released in estuaries; the subsequent zoeal-​stage larvae developed in open coastal waters and



Larval Growth and Development

showed a reduced capacity in osmoregulation. The osmoregulatory capacity increased again during the last zoea stage (zoea IV) and megalopal stage, likely to prepare for the megalopae to recolonize estuaries and coastal lagoons. The reduced capacity to osmoregulate in zoea II and III larvae living in open waters as seen in N. granulata and other species may represent a strategy to reduce the costs of osmoregulation and thus improve growth (Charmantier et al. 2002). Similarly, in the sesarmid crab Armases roberti, zoea I larvae hatch in rivers and show a weak osmoregulatory ability; these larvae are presumably exported quickly to coastal waters with higher salinities. By the megalopal stage, they become more tolerant to low salinity and migrate back to freshwater habitats to settle (Torres et al. 2006). In fact, a similar pattern of greater osmoregulatory strength at the megalopal stage was found in various crabs and is believed to be linked to better developed gills with enhanced Na+/​K+-​ATPase activity (Cieluch et al. 2004, 2007). In the mud crab Scylla serrata, a laboratory experiment showed relatively little change in salinity tolerance through consecutive zoeal instars (Nurdiani and Zeng 2007). This is probably linked to the fact that despite S. serrata inhabiting estuarine and coastal waters, the mature females migrate long distances offshore to release their larvae, where zoeae complete their development. A similar situation was found for oceanic and deep-​water dwelling species, such as the snow crab Chionoecetes opilio (Yamamoto et al. 2015). In fact, although many estuarine crustacean species are considered euryhaline and osmoregulating, a large number of them export their larvae to the ocean, and osmoregulation mainly occurs during juvenile and adult stages (also, probably to a certain extent, at postlarval stages; e.g., megalopa). During their larval development in the sea, they are similar to those of marine species and are stenohaline osmoconformers. Osmotic stress could drastically affect the growth and biochemical composition of crustacean larvae, which often occurs as a reduction in dry mass, lipids, and/​or protein content (Torres et al. 2002). This could be a result of an impairment or readjustment of larval metabolism, but may also be a consequence of reduced feeding activities under osmotic stress (Anger 2003). In general, however, the biochemical composition of stronger osmoregulators is not as severely affected as that of osmoconformers (Fig. 7.3) (Torres et al. 2002, Anger 2003, Torres et al. 2011). Even for late embryos, exposure to suboptimal salinities could negatively influence the hatching success or biomass of newly hatched larvae, as seen in the calanoid copepod Acartia tonsa (Peck and Holste 2006) and the estuarine crab N. granulata (Giménez and Torres 2002). A likely explanation for this could be that suboptimal salinities reduced nutrient reserves in the eggs because they were used for osmoregulation by the embryos (Charmantier and Charmantier-​Daures 2001). Moreover, exposing zoeal larvae of N. granulata to a lower salinity was also found to increase the frequency of the larvae undergoing an extra instar (i.e., five rather than four zoeal instars) before reaching the megalopal stage (Giménez and Torres 2002). It is likely that the need to osmoregulate under such low salinities requires extra time during zoeal development to accumulate sufficient reserves for successful metamorphosis (Giménez and Torres 2002). Interestingly, a short-​term exposure of the spider crab H.  araneus first-​stage zoeae to low salinity (15) not only caused delayed development and reduced biomass at the zoea 1 stage, but also led to delayed effects at subsequent larval stages (Anger 1985). Moreover, suboptimal salinities were observed to induce morphological alterations in larvae of the crabs Rhithropanopeus harrisii (Goncalves et al. 1995) and Eriocheir sinensis (Anger 2003). In the latter case, larval carapace spines were found to be proportionally longer (in relation to body length) and increased during larval development at low salinities. Anger (2003) suggested that this might be an adaptive response to increase buoyancy at low salinities and thus conserve energy for position maintenance in the water column. Such a response was suggested as particularly important because larval reserves become increasingly depleted under osmotically stressful conditions. Plasticity in salinity tolerance has also been shown in larval decapods. Experiments indicated that salinity experienced during the embryonic phase could influence osmotic tolerance of the

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Fig. 7.3. Increased capacity to hyperosmoregulate reduces or eliminates the negative effects of low salinity on accumulation of biomass (dry mass, proteins, and lipids) in decapod larvae. The graphs show the negative correlation between the osmoregulatory capacity (OC) and the biomass-​specific effect of salinity 20 (B20ʹ); all correlations are significant except for proteins versus OC20. OC is the difference between hemolymph osmolality and the osmolality expected in case of complete osmoconformation, which is here calculated at salinity 20 (OC20)—​the minimal salinity for which survival was more than 80% (OCM). B20ʹ is the relative difference in biomass between larvae kept in seawater and at salinity 20, per unit of salinity. Species and stages are as follows: 1 through 4, Armases miersii (zoea I–​III, megalopa); 5 through 9, Armases roberti (zoea I–​IV, megalopa); 10 through 14, Armases ricordi (zoea I–​IV, megalopa); 15 through 22, all zoea I as follows: 15, Sesarma curacaoense; 16, Perisesarma fasciatum; 17, Neohelice granulata; 18, Carcinus maenas; 19, Hyas araneus; 20, Homarus gammarus; 21, Nephrops norvegicus; and 22, Cancer pagurus. Circles mark the most osmoconforming stages. Modified from Torres et al. (2011), with permission from © Inter-​Research 2011.

subsequent larvae (Rosenberg and Costlow 1979, Laughlin and French 1989b; also see Chapter 10 in this volume). In addition, acclimation could be crucial for the salinity tolerance of advanced larval stages. For example, gradual acclimation reportedly allowed A. roberti megalopae to tolerate fresh water whereas an abrupt transfer from salinity of 25 to ≤3 caused total mortality within 24 hours (Torres et al. 2006). It is worth noting, however, that adult estuarine copepods (E. affinis)



Larval Growth and Development

acclimated to low salinities did not lead to offspring capable of coping better with low salinities (Lee and Petersen 2003). Nevertheless, postembryonic E. affinis showed higher individual variations in their salinity tolerance (Devreker et al. 2007), leading to the suggestion that such a characteristic facilitates invasion of the species into freshwater environments (Lee and Peterson 2003, Lee et al. 2003).

OTHER PHYSICOCHEMICAL FACTORS Hypoxia Most crustacean larvae develop in well-​oxygenated waters and the rate of larval oxygen consumption or respiration is highly correlated with the overall rates of metabolic processes. Nevertheless, crustacean larvae may experience hypoxic conditions, for example, at the time of larval release (Yannicelli et al. 2013), when sinking to deeper waters during diel vertical migration (Spicer and Stromberg 2003), or during settlement (Tankersley and Wieber 2000). Larval hemocyanins generally have lower oxygen affinity than those of juveniles and adults [reviewed in Spicer (1995)] and hence they may be less tolerant to hypoxia. Low oxygen levels are known to decrease respiration rates in decapod larvae (Anger 2001), although some species are more adaptive to such conditions. For instance, both zoeae and megalopae of the bromeliad crab Metopaulias depressus could tolerate dissolved oxygen as low as 14% to 21% saturation, which is believed to be an adaptive trait because larval development of this species occurs in small puddles of rainwater collected at the base of bromeliad plants, where oxygen levels could be low (Diesel and Schuh 1993). During episodes of oxygen depletion, larvae can encounter environmental and functional hypoxia, which are both physiologically stressful. In laboratory studies, it has been shown that in crustaceans, the ability to respond to hypoxia can change during larval ontogeny (Tankersley and Wieber 2000, Spicer and Eriksson 2003, Spicer and Strömberg 2003, Yannicelli et  al. 2013, Alter et al. 2015). For example, euphausids pass through a complex larval phase comprised of nauplius, metanauplius, calytopis, and furcilia (Olesen 2018); in the northern krill Meganyctiphanes norvegica, early larvae remain in shallow waters but later larvae (postfurcilia VI) perform diel vertical migration during which hypoxic waters are encountered. A study by Spicer and Stromberg (2003) found that the capacity to oxyregulate appeared at furcilia III and became stronger at furcilia V, when the larvae were able to hyperventilate. Exposure to hypoxia reportedly did not lead to significant differences in size or biomass in the larvae of the crabs Petrolisthes laevigatus and Lithodes santolla (Alter et  al. 2015); nor were there significant differences found in the biomass or altercation of the allometric growth in the larval and early juvenile Norway lobster Nephrops norvegicus (Spicer and Eriksson 2003). However, in the squat lobster Pleuroncodes monodon, which inhabits depths of 150 to 200 m with low dissolved oxygen, a study found that its zoeae survived hypoxia but were oxyconforming (i.e., their respiration was depressed, especially in zoea I); however, the megalopae were oxyregulating. Although the zoea I could still grow under hypoxia, the accumulation of carbon and nitrogen was reduced, and development was delayed considerably (Yannicelli et al. 2013). In the blue crab C. sapidus, recruiting to estuaries could expose megalopae to summer hypoxia, and it was found that metamorphosis of megalopae to the first crabs was delayed when they were exposed to low oxygen concentrations (Tankersley and Wieber 2000). Similarly, N. norvegicus may experience hypoxia during larval settlement; preexposure of larval N. norvegicus to low oxygen levels led to a shift from oxyconformation to oxyregulation, pointing to the existence of adaptive plasticity in larval response to hypoxia (Spicer and Eriksson 2003).

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Developmental Biology and Larval Ecology Light Light is a complex, highly variable environmental parameter that serves as both taxis and kinesis stimulus to crustacean larvae (Sulkin 1984; see Chapter 12 in this volume). In aquatic environments, there are four important aspects of underwater light affecting crustacean larvae:  intensity, spectral composition, angular distribution, and polarization. The vast majority of crustacean larvae are planktonic and live in a light field of open waters, which represents a relatively simple and predictable photic environment. The relative basic visual requirements of these larvae, and the need to remain transparent to reduce predation risk, lead to the use of a single eye type by marine crustacean larvae. This is opposed to very complicated and diversified visual systems of adult crustaceans that occupy highly diverse habitats (Cronin and Jinks 2001, Cronin and Feller 2014, Glantz 2014). It is believed that the vision of crustacean larvae is primarily concerned with orientation in the water column, vertical migration, and avoidance of predators, rather than feeding as in fish larvae, while the bulk of research in this area comes from studies on brachyuran crab larvae (Forward 1976, 1986, Sulkin 1984, Forward and Buswell 1989, Epifanio and Cohen 2016). Absorption and scattering lead to decreasing light intensity with water depth (Epifanio and Cohen 2016). However, there is limited information on the effects of light intensity on survival, development, and growth of crustacean larvae, and a few such studies are conducted in the laboratory for aquaculture purposes. For instance, it was found that light intensity did not significantly affect growth and survival of early phyllosoma larvae of the rock lobster Jasus edwardsii (Moss et al. 1999). Similarly, a more recent study of the American lobster Homarus americanus reported no significant effects of light intensity on both larval survival and growth, although significant interaction between light intensity and photoperiod on larval survival was detected (Haché et al. 2015). Crustacean larvae respond to a fairly broad range of wavelengths, although response peaks vary among species and through ontogeny (Sulkin 1984, Hamasaki et al. 2013, Johnson and Rhyne 2015). It is worth noting that crustacean larvae are considered vulnerable to solar ultraviolet (UV) radiation, because many of them live in the upper layers of the water column and are generally not well pigmented to reduce the risk of being preyed upon by visual predators. Increased mortality caused by UV radiation has been reported in late copepodites of calanoid copepods (Ban et al. 2007) and crab zoeae (Al-​Aidaroos et al. 2015). Some crustacean larvae are filter feeders and light is not required for their feeding. Light is also not required for prey capture in carnivorous crustacean larvae. Feeding by carnivorous crustacean larvae is believed to rely on random encounters and/​or olfactory cues (Anger 2001), and it occurs in darkness. However, there were reports suggesting that feeding rates of crab zoeae during the light phase were several times higher than those during the nighttime (Yatsuzuka 1962, Minagawa and Murano1993). Coincidently, recent laboratory experiments have demonstrated that photoperiod, which is normally associated with seasonal changes in the wild but often manipulated in laboratory and aquaculture settings, could significantly affect the growth and development of some crustacean larvae. Such effects were generally attributed to light-​induced changes in larval swimming/​ feeding activity. For example, larval development of the calanoid copepod Acartia sinjiensis was significantly affected by photoperiod, with a clear trend of development accelerated with increasing illumination duration (Camus and Zeng 2008). Similarly, in the blue swimmer crab P. pelagicus, both the survival and development of zoeal larvae to megalopae were affected by photoperiod, with zoeal development significantly faster under photoperiods with a longer light phase (18 hours and 24 hours of light) than under constant darkness. In addition, newly molted megalopae raised in constant darkness had the smallest carapace length and dry weight (Andrés et al. 2010). Similarly, in the Australian giant crab Pseudocarcinus gigas, longer photoperiods and brighter light were found to lead to shorter intermolt durations, with the most rapid development to megalopa found under continuous light; the smallest late zoeae (zoea IV) were also found in the continuous dark treatment



Larval Growth and Development

(Gardner and Maguire 1998). Photoperiod manipulations also significantly affected growth, development, and feeding of larvae of the spiny lobsters J.  edwardsii (Bermudes and Ritar 2008) and Sagmariasus verreauxi (Fitzgibbon and Battaglene 2012) with a shift in optimal photoperiod during larval ontogeny. Increased Carbon Dioxide Levels Oceans have absorbed more than one third of atmospheric carbon dioxide since the beginning of the Industrial Revolution, leading to a 0.1-​U decrease in pH and ~16% reduction in carbonate ion concentration. By the end of the 21st century, pH in the world oceans are predicted to drop from the current ~8.1 to ~7.8 [reviewed by Gatusso and Hansson (2011)]. Marine organisms producing calcified shells are a research focus on ocean acidification (OA) because the formation of such structures are affected by high partial pressure of carbon dioxide (pCO2), low pH, and low CO2–​ (e.g., Byrne 2011, Przelawski et al. 2015). Although crustaceans have a mineralized exoskeleton, they have received comparatively less attention than mollusks in such studies, and even more so for their larvae (Whiteley 2011). A key aspect defining the tolerance of crustaceans to OA is their capacity for osmo-​and ion regulation, which also defines their capacity to compensate acid–​base disturbances produced by OA (Fig. 7.4A). As a consequence of ion regulation (Fig. 7.4B), bicarbonate ion (HCO3–​) in hemolymph could be higher and such crustaceans appear to tolerate hypercapnia better. Hence, vulnerability may be higher for osmoconformers, which is most common among crustacean larvae (see the “Salinity” section in this chapter). Three characteristics of crustacean larvae distinguish them from other life history stages. First, larval exoskeleton has little to no calcification, although some calcification does occur during postlarval stages in some decapods (e.g., megalopae of crabs) (Anger 2001). Second, larvae of most crustaceans are planktonic and tend to be active swimmers. Third, osmoregulation capacity could vary substantially among species and during larval ontogeny (see the “Salinity” section in this chapter). Hence, variable responses of different crustacean larvae and larval stages to increasing pCO2 are expected. Walther et al. (2010, 2011) and Schiffer et al. (2013) evaluated the effect of increased pCO2 on the larval survival, development, and growth of the marine crab H. araneus, an osmoconformer. They found that exposure to high pCO2 levels resulted in extended larval development and reduced larval calcium contents in both populations from Svalbard and Helgoland, the northernmost (coldest) and the southernmost (warmest) end of the species’ distribution range, respectively. However, the reduction in dry weight and carbon-​to-​nitrogen ratio (as a proxy of the lipid content) was significant only in larvae originating from Helgoland. Similarly, effects of elevated pCO2 levels on reducing dry weight, calcium, and magnesium were reported in the last larval stage of Homarus gammarus (Arnold et al. 2009). However, in a study of larvae of the barnacle Amphibalanus improvisus, it was found larval development or size was not significantly affected by the reduced pH, leading to the suggestion that the noncalcifying larval stages of A. improvisus are generally tolerant to near-​future levels of OA. This finding is in line with the reports of other barnacles, suggesting barnacles do not show a greater sensitivity to OA in early life history as reported for other invertebrates (Pansch et al. 2013). On the other hand, when subjected to a combination of elevated pCO2 and temperature, although no significant difference in larval survival of A. improvisus was reported (Pansch et al. 2013), increased mortality was observed for larvae of another barnacle species: Balanus amphitrite (Baragi and Anil 2015). Interestingly, when embryos of Hyas araneus were exposed to OA conditions, stronger effects led to increased larval mortalities, delay in larval development, and decreases in growth in terms of carbon content and dry weight, as well as reduced larval feeding rates (Schiffer et  al. 2014a).

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This suggests that exposure to OA conditions during embryogenesis likely produces physiological perturbations instead of adaptive plasticity in the larvae. In summary, although efforts have been made to quantify OA effects on crustacean larvae, more research is needed to understand more fully the potential adaptive mechanisms that may help them to survive in increasingly acidified habitats. To evaluate the hypothesis that ion regulation ability may represent a preadaptation to low pH, further experiments should be conducted on larvae that undergo development in estuaries or tidal pools because such larvae generally show stronger osmoregulation capacities (Anger et al. 2008). Pollutants Among various pollutants present in aquatic environments, heavy metals, insecticides, and crude oil have traditionally received more attention on their toxicity to crustacean larvae (e.g., Rainbow 1995, Ahearn et al. 2004, Osterberg et al. 2012, Sun et al. 2014). Recently, however, there are more studies evaluating the effects of emergent contaminants such as pharmaceutical compounds,



Larval Growth and Development

microplastics (MPs), and plasticizers on crustacean larvae (e.g., Gonzalez-​Ortegón et  al. 2013, Heindler et al. 2017). Overall, it is a common phenomenon that crustacean larvae exhibit markedly higher sensitivity to various pollutants than juveniles and adults (e.g., Wong et al. 1995, Forget-​Leray et al. 2005, Kulkarni et al. 2013, Gomes et al. 2016). The toxicity of heavy metals to crustacean larvae has been studied for decades with a focus on lead (Pb), mercury (Hg), cadmium (Cd), copper (Cu), and zinc (Zn) (Rainbow 1995). Toxicity studies are usually first based on acute tests to estimate LC50—​the concentration causing 50% mortality of the tested population within a given exposure period. Based on LC50, heavy-​metal acute toxicity to crustacean larvae is often species specific. For instance, although Cd was found more toxic than Hg or Cu to the first-​stage larvae of the spider crab M.  brachydactyla and the lobster H. gammarus, Hg was more than 20-​fold more toxic than Cu and Cd to the first-​stage larvae of P.  serratus (Mariño-​Balsa et  al. 2000). Meanwhile, sublethal exposure to lower concentrations of heavy metals reportedly led to delayed development and reduced growth, partially as a result of decreased feeding activity. Because larvae with delayed development spend more time in the water column, they are also more susceptible to predation in the wild (Wong et  al. 1993, 1995). Heavy-​metal exposure may also induce deformities in larvae (Amin et al. 1998, Shealy and Sandifer 1975). Embryos of the king crab Lithodes santolla exposed to elevated Pb and Cd levels led to newly hatched larvae with various deformities, including atrophy of the dorsal, rostral, and telson spines, as well as pereopods and telson setae (Amin et al. 1998). Similarly, during larval development of the shrimp Palaemonetes pugio, exposure to sublethal levels of Hg led to additional instars along with increased morphological variability and deformities (Shealy and Sandifer 1975). In general, larval tolerance to heavy metals increases with development, likely a result of improved detoxification mechanisms (Wong et al. 1995). Among these, metallothionein (MT), a protein present in animals that binds heavy metals, is known to reduce heavy-​metal toxicity. The role of MT in crustacean larvae was first confirmed in the crab R. harrisii exposed to Cu (Sanders et al. 1983). More recently, Sun et al. (2014) reported that the expression of MT-​I increased in zoea I and megalopae of the mitten crab E. sinenesis exposed to Cu. In fact, MT has been suggested to be responsible for increased tolerance to Cd, Cu, and Zn by zoeae of Metacarcinus anthonyi hatched from embryos exposed to them (Macdonald et al. 1988). Crustacean larvae can be particularly sensitive to insecticides because their modes of toxicity by disrupting or inhibiting key biological pathways (e.g., neurotransmission or endocrine system) are often shared between insects and crustaceans (Rodríguez et al. 2007). Insecticide use is widespread in agriculture and, during heavy rains, the runoff enters aquatic environments, which often form important habitats for crustacean larvae. Crustacean larvae generally are more sensitive to insecticides than juveniles or adults (e.g., Osterberg et al. 2012). Various insecticides, such as heptachlor (endocrine disrupter), glyphosate (herbicide), and pyriproxyfen, fenoxycarb, and methoprene (juvenile hormone analogues), have been reported to increase mortality, delay molting, and induce abnormalities in various decapod larvae (Hertz and Chang 1986, McKenney and Celestial 1993, Snyder and Mulder 2001, Tuberty and McKenney 2005, Avigliano et al. 2014). For example, methoprene at concentrations ≥8 μg/​L was found to inhibit metamorphosis and increase respiration throughout the larval phases of the shrimp P. pugio, but premetamorphic larvae showed a growth acceleration, possibly resulting from increased lipid catabolism (McKenney and Celestial 1993). Nitrogenous wastes including ammonia, nitrite, and nitrate are a global issue in aquatic ecosystems as a result of escalating anthropogenic activities leading to eutrophication. The relative toxicity of these inorganic pollutants to crustacean larvae is similar to adults (i.e., ­ammonia  >  ­nitrite  > nitrate-​nitrogen) (Romano and Zeng 2013), although larvae are usually much more sensitive. For example, nauplii of the copepod Parvocalanus crassirostris were found to be eight times more sensitive to ammonia exposure than adults, with a 48-​hour LC50 at only 1.28  mg/L. Moreover, among a range of biological parameters tested, larval development was

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Developmental Biology and Larval Ecology found to be most sensitive to sublethal ammonia exposure, with significant delay detected at a concentration as low as 0.06 mg/​L. This suggests that, even with ammonia levels currently found in coastal waters, the sublethal effects could potentially influence population dynamics of the species, highlighting harmful effects of effluent disposal to coastal ecosystems (Middleton 2013). Unlike most other pollutants, crude oil remains on the surface waters, potentially leading to additional physical disruptions to the larvae. Crude oil also contains various compounds, including polycyclic aromatic hydrocarbons (PAHs), which are lipophilic and thus can accumulate in tissues (Ellis et al. 1995). Under UV radiation, PAHs can potentially become more toxic; such photo-​induced toxicity was indeed reported to affect larvae of various crabs, including Libinia dubia, Menippe dina, Panopeus herbstii (Peachey 2005), and C. sapidus (Alloy et al. 2015). Photo-​ induced toxicity is believed to be particularly relevant to larvae of benthic decapods because they often distribute high in the water column and are therefore exposed to high UV radiation (Peachey 2005). Increased rates of protein catabolism, delayed molting, and reduced growth were reported among crude oil-​exposed American lobster H. americanus larvae, attributed to disruptions of lipid metabolism possibly as a detoxifying response (Capuzzo et al. 1984). However, although sublethal levels of naphthalene (45–​300 μg/​L), the simplest PAH, were found to increase respiration rates in the larval mud crab R. harrisii, the effects declined with larval ontogeny, and when growth was measured at the megalopal stage, no significant effect was detected (Laughlin and Neff 1981). The toxicity of PAHs might be reduced by dispersants designed to emulsify and facilitate oil degradation (Alloy et al. 2015). However, at the concentrations used to treat oil spills, dispersants have been found to affect swimming activity of zoeae of C. sapidus and nauplii of various copepods, including A.  tonsa, Temora turbinate, and Pseudodiaptomus pelagicus (Pie and Mitchelmore 2015, Almeda et al. 2016). Moreover, Almeda et al. (2016) found that oil emulsified by the dispersant was actually more toxic than crude oil to copepod nauplii. Recently, effects of MPs (size,