Miscellaneous Invertebrates 3110488337, 9783110488333

Table of contents :
Preface
Contents
List of contributing authors
1. Dicyemida
2. Orthonectida
3. Placozoa
4. Seisonidae
5. Cycliophora
6. Entoprocta (Kamptozoa)
7. Chaetognatha
8. Pterobranchia
9. Enteropneusta
Index

Citation preview

Handbook of Zoology Miscellaneous Invertebrates

Handbook of Zoology Founded by Willy Kükenthal continued by M. Beier, M. Fischer, J.-G. Helmcke, D. Starck and H. Wermuth Editor-in-chief Andreas Schmidt-Rhaesa

Miscellaneous Invertebrates Edited by Andreas Schmidt-Rhaesa

DE GRUYTER

Miscellaneous Invertebrates

Edited by Andreas Schmidt-Rhaesa

DE GRUYTER

Scientific Editor Andreas Schmidt-Rhaesa University of Hamburg Martin-Luther-King-Platz 3 20146 Hamburg, Germany

ISBN 978-3-11-048833-3 e-ISBN (PDF) 978-3-11-048927-9 e-ISBN (EPUB) 978-3-11-048841-8 ISSN 2193-2824 Library of Congress Control Number: 2018951344 Bibliografic information published by the Deutsche Nationalbibliothek The Deutsche Nationalbibliothek lists this publication in the Deutsche Nationalbibliografie; detailed bibliographic data are available in the Internet at http://dnb.dnb.de. © 2019 Walter de Gruyter GmbH & Co. KG, Berlin/Boston Typesetting: Compuscript Ltd. Shannon, Ireland Printing and Binding: CPI books GmbH, Leck www.degruyter.com

Preface This volume of the series Handbook of Zoology includes animal taxa that are, roughly spoken, too small to fill their own volume and too separate from other taxa to be included in a volume on another group. Some of them are phylogenetically still not settled. Dicyemida and Orthonectida, often united as “Mesozoa”, are still mysterious in many aspects and not settled among the non-bilaterian metazoans. Placozoa also enjoy different phylogenetic positions in the basal region of the animal tree. Seisonidae belong into Gnathifera, close to rotifers and acanthocephalans. They should have been included in the “Gastrotricha and Gnathifera” volume previously published but had to be delayed to this volume. Cycliophora, described in 1995, are protostomes with a fascinating life cycle and a still not certain phylogenetic position.

https://doi.org/10.1515/9783110489279-202

Kamptozoa or Entoprocta are trochozoans, but their small size in terms of species number and available literature compared to the other large trochozoan groups such as annelids and molluscs justifies their inclusion into this volume. Chaetognatha have puzzled systematists since forever, and although the circles are getting narrower (they seem to be protostomes), the exact position is still uncertain. Finally, Enteropneusta and Pterobranchia, the hemichordates, are very likely the sister group of Echinodermata (together Ambulacraria). I am very thankful to all authors who have contributed to this wonderful volume, which hopefully will serve as an up to date review for these groups. Andreas Schmidt-Rhaesa

Contents Preface

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List of contributing authors

xi

Hidetaka Furuya 1 Dicyemida 1 1.1 Introduction 1 Life cycle 1.2 1 Body organization 1.3 2 Species diversity of dicyemids 1.4 2 Development of embryos 1.5 3 Infusoriform embryos 1.5.1 3 Vermiform embryos 1.5.2 4 1.6 Gametogenesis in hermaphroditic gonad 4 1.7 Reproductive strategy in the renal organs 7 Literature 8 George S. Slyusarev 2 Orthonectida 11 2.1 Introduction 11 2.2 Morphology of free-living females and males 11 General and external morphology 2.2.1 11 The structure of epithelial cells 2.2.2 13 2.2.3 The structure of the cuticle in free-swimming (mature) adults 14 Formation of the cuticle 2.2.4 14 2.2.5 Cilia 18 Cytoskeleton of epithelial cells 2.2.6 20 The cytoplasm of the ciliated cells 2.2.7 20 2.2.8 The genital pore in a non-fertilized female 21 The genital pore in a fertilized female 2.2.9 21 2.2.10 Musculature 21 The nervous system 2.2.11 26 Ciliated receptor 2.2.12 27 2.2.13 Gametes 27 Cell number counts 2.2.14 29 2.3 Plasmodium 29 Plasmodial cytoplasm 2.3.1 30 Reproductive cells 2.3.2 30 Plasmodial feeding 2.3.3 31 2.3.4 Egress of males and females from the plasmodium 33 2.3.5 Plasmodial reproduction 33

2.3.6 The nature of the orthonectid plasmodium 34 Reproduction and development 2.4 35 Reproductive biology 2.4.1 35 2.4.2 Embryology 35 2.4.3 Larva 35 2.5 Systematics 35 2.6 Distribution, biology, and ecology 36 The life cycle 2.7 37 Acknowledgments 37 Literature 38 Oliver Voigt and Michael Eitel 3 Placozoa 41 3.1 Introduction 41 3.2 Distribution 41 3.3 Biology 43 3.3.1 Anatomy 43 Upper epithelium 3.3.1.1 43 Lower epithelium 3.3.1.2 43 Intermediate layer 3.3.1.3 43 Movement and feeding 3.3.2 45 Asexual reproduction 3.3.3 46 3.3.4 Sexual reproduction and early embryonic development 47 3.3.5 Ecology 47 Phylogeny and taxonomy 3.4 49 Placozoa in the animal tree of life 3.4.1 49 Genetic diversity and phylogeography 3.4.2 49 3.4.3 Genomics 51 Literature 51 Wilko H. Ahlrichs and O. Riemann 4 Seisonidae 55 4.1 Introduction 55 4.2 Morphology 55 General and external morphology 4.2.1 Body outline 4.2.1.1 55 4.2.1.2 Head 56 4.2.1.3 Neck 58 4.2.1.4 Trunk 58 4.2.1.5 Foot 58 4.2.2 Epidermis 59 Epithelium of the epidermis 4.2.2.1 59 4.2.2.2 Epidermal glands 61 4.2.3 Musculature 67 4.2.3.1 Circular muscles 67

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4.2.3.2 Longitudinal muscles (body retractors) 68 Longitudinal muscles of P. annulatus 4.2.3.3 68 Longitudinal muscles of S. nebaliae 4.2.3.4 68 4.2.3.5 Ultrastructure of the longitudinal muscles (see Ahlrichs 1995, Ricci et al. 1993) 68 Nervous system 4.2.4 68 4.2.4.1 Cerebral ganglion 68 Sensory structures 4.2.5 69 Receptor of the rotatory organ 4.2.5.1 69 Stretch receptors 4.2.5.2 69 Stomach receptor 4.2.5.3 69 Dorsal antenna 4.2.5.4 69 Lateral head antennae 4.2.5.5 69 Foot plate receptor 4.2.5.6 69 Mastax receptor 4.2.5.7 69 Intestinal system 4.2.6 69 4.2.6.1 Mouth 69 4.2.6.2 Esophagus 70 4.2.6.3 Ultrastructure 70 4.2.6.4 Stomach 70 4.2.6.5 Cloaca 70 4.2.6.6 Feeding and diet 70 4.2.6.7 Mastax 70 4.2.6.8 Terminology 72 4.2.6.9 General mastax organization 72 4.2.6.10 Incus 72 4.2.6.11 Epipharynx 72 4.2.6.12 Hypopharynx 72 4.2.6.13 Malleus 73 4.2.6.14 Ultrastructure of the mastax 73 4.2.6.15 General ultrastructure of trophi 73 4.2.6.16 Ultrastructure of trophi of Seisonidae (see Ahlrichs 1995) 73 4.2.6.17 Mastax muscles 74 4.2.7 Body cavity 74 4.2.8 Excretory system 75 4.2.8.1 Terminal syncytium 76 4.2.8.2 Canal syncytium 76 4.2.8.3 Multiciliated canal region 77 4.2.8.4 Main canal 77 4.2.8.5 Nephroporus cell 77 4.2.9 Reproductive organs 77 4.2.9.1 Female gonad 77 4.2.9.2 Male gonad 77 4.2.10 Gametes 79 4.2.10.1 Spermatogenesis and spermiogenesis 79 4.2.10.2 Oogenesis 80 4.2.10.3 Fertilization 81 4.2.10.4 Fertilized eggs 82 4.3 Cleavage and development 82 4.4 Ecology and Physiology 82

82 4.4.1 Habitat Oxygen requirements 4.4.2 82 Seasonal occurrence 4.4.3 83 4.4.4 Movements 83 4.5 Phylogeny 83 4.6 Systematics 83 Hints on identification 4.6.1 83 4.7 Biogeography 84 Acknowledgments 84 Literature 84 Peter Funch and Ricardo Neves 5 Cycliophora 87 5.1 Introduction 87 Life cycle 5.2 87 5.3 Morphology 91 General morphology 5.3.1 91 5.3.2 Integument 91 5.3.3 Musculature 95 5.3.4 Nervous system 98 5.3.5 Sensory structures 101 5.3.6 Digestive tract 102 5.3.7 Connective tissue and body cavities 102 5.3.8 Excretory system 102 5.3.9 Reproductive organs 103 5.3.10 Gametes 104 5.4 Reproduction and development 104 5.4.1 Reproductive biology 104 5.4.2 Cleavage and development 107 5.5 Phylogeny 107 5.6 Diversity 108 5.7 Ecology 108 Acknowledgments 109 Literature 109 Anastasia O. Borisanova 6 Entoprocta (Kamptozoa) 111 6.1 Introduction 111 6.2 General morphology 111 6.3 Integument 114 6.3.1 Cuticle structure 114 6.3.2 Epidermis 115 6.3.3 Glands 115 6.4 Musculature 118 6.5 Nervous system 123 6.5.1 Sense organs 124 6.6 Digestive system 125 6.6.1 Food transport 128 6.7 Tentacle structure and filter-feeding mechanism 128 6.8 Body cavity 131

Contents 

133 6.9 Excretory system 6.10 Regeneration 134 Reproductive system 6.11 136 6.11.1 Oogenesis 136 6.11.2 Spermatogenesis 139 6.12 Fertilization and embryogenesis 140 Larval morphology 6.13 142 6.13.1 Introduction 142 6.13.2 Integument 143 Digestive system 6.13.3 144 Excretory system 6.13.4 145 6.13.5 Musculature 145 Nervous system 6.13.6 146 Apical organ 6.13.7 146 Frontal organ 6.13.8 146 6.13.9 Metamorphosis 147 6.14 Paleontology 148 6.15 Phylogeny 149 6.16 Biogeography and ecology 150 6.16.1 Biogeography 150 6.16.2 Ecology 151 6.17 Systematics 152 Literature 157 Carsten H.G. Müller, Steffen Harzsch and Yvan Perez 7 Chaetognatha 163 7.1 Introduction 163 7.1.1 Generalities 163 7.1.2 Paleontological evidence 165 7.1.3 Gross morphology and anatomy 166 7.1.4 Affinities of Chaetognatha within Bilateria 170 7.2 Systematics and taxonomy 173 7.2.1 Traditional hypotheses based on morphology 173 7.2.2 Evidence from recent molecular phylogenies 175 7.2.3 Ecophenotypic plasticity 177 7.2.4 Species diagnosis, DNA bar coding, and population genetics 178 7.2.5 List of valid species by family and genus 179 7.3 Ecology and distribution patterns 185 7.3.1 Species composition and distribution in the major ocean basins 185 7.3.1.1 Planktonic species 189 7.3.1.2 Benthic species 192 7.3.1.3 Endemism and rarely recorded species 193 7.3.2 Diel and ontogenetic vertical migrations 194

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7.4 Histological structure, cytology, and functional significance of organ systems 196 7.4.1 Epidermis 197 7.4.1.1 Distal epidermal cells 197 7.4.1.2 Proximal epidermal cells 197 7.4.1.3 Vacuolated epidermal cells 199 7.4.1.4 Further types of epidermal cells 203 7.4.1.5 Oligo- and unilayered epidermis and chitinous structures 203 7.4.1.6 Vestibular organs 205 7.4.1.7 Multicellular epidermal glands and solitary secretory cells 207 Bioluminescent organs 7.4.1.8 211 7.4.2 Sense organs 212 7.4.2.1 Retrocerebral organ 212 7.4.2.2 Eyes 213 7.4.2.3 Ciliary fence and tuft organs 216 7.4.2.4 Corona ciliata 219 7.4.3 Nervous system 221 7.4.3.1 Brain and circumesophageal chain of cephalic ganglia 223 7.4.3.2 Ventral nerve center 227 7.4.3.3 Intra- and basiepidermal neuronal plexus 229 7.4.4 Muscular apparatus and locomotion 231 7.4.4.1 Gut-associated muscles 233 7.4.4.2 Cephalic locomotory muscles 233 7.4.4.3 Trunk and tail locomotory muscles 235 7.4.4.4 Neuromuscular systems 240 7.4.4.5 Considerations about muscle evolution within the Chaetognatha 241 7.4.5 Digestive system 242 7.4.5.1 Oral cavity and esophagus 242 7.4.5.2 Intestine and rectum 242 7.4.5.3 Vacuolated intestine 245 7.4.6 Body cavities 245 7.4.6.1 Primary body cavity—hemal system 245 7.4.6.2 Secondary body cavities—trimeric coelomic organisation 247 7.4.6.3 Status and phylogenetic significance of chaetognath coeloms and associated metanephridial organs 249 7.4.7 Reproductive system 251 7.4.7.1 Testis and seminal vesicle 253 7.4.7.2 Ovary and oviducal complex 255 7.5 Reproduction and ontogeny 256 7.5.1 Mating, fertilization, and egg laying 256 7.5.2 Germline, cleavage, gastrulation, and coelomogensis 261

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7.5.3 Hatching and growth 7.5.4 Neurogenesis 265 Literature 267

265

Kenneth M. Halanych, Michael G. Tassia and Johanna T. Cannon 8 Pterobranchia 283 8.1 Introduction 283 8.2 Morphology 283 General anatomy 8.2.1 283 8.2.2 Integument 286 Body cavities 8.2.3 287 8.2.4 Musculature 287 8.2.5 Nervous system and sensory structures 287 Digestive system 8.2.6 288 Gas exchange system 8.2.7 288 Excretory and circulatory 8.2.8 systems 289 8.3 Reproduction and development 289 8.3.1 Reproductive system 289 8.3.2 Development 290 8.4 Distribution and ecology 292 8.5 Phylogeny 292 8.6 Diversity 293 Literature 296

Michael G. Tassia, Johanna T. Cannon and Kenneth M. Halanych 9 Enteropneusta 299 9.1 Introduction 299 9.2 Morphology 299 General anatomy 9.2.1 299 9.2.2 Integument 303 Body cavities 9.2.3 304 9.2.4 Musculature 305 9.2.5 Nervous system and sensory structures 307 Digestive and Branchial 9.2.6 systems 309 Digestive system 9.2.6.1 309 Branchial system 9.2.6.2 310 Excretory and circulatory 9.2.7 systems 311 Reproduction and development 9.3 312 9.3.1 Reproductive system 312 9.3.2 Development 312 9.4 Distribution and ecology 315 9.5 Phylogeny 315 9.6 Diversity 316 Literature 321 Index 

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List of contributing authors Wilko Ahlrichs E-mail: [email protected] Institute for Biology und Environmental Sciences (IBU) University of Oldenburg PO-Box 2503, Oldenburg 26111, Germany Anastasia O. Borisanova E-mail: [email protected] Department of Invertebrates Zoology Faculty of Biology Moscow State University 1-12 Leninskie Gory, Moscow 119991, Russia. Johanna T. Cannon E-mail: [email protected] Department of Ecology, Evolution, and Marine Biology University of California Santa Barbara Santa Barbara, CA 93106, USA Michael Eitel E-mail: [email protected] Department of Earth and Environmental Sciences Ludwig-Maximilians-University of Munich Richard-Wagner-Str. 10, Munich 80333, Germany Peter Funch E-mail: [email protected] Department of Bioscience Aarhus University Ny Munkegade 116, building 1540, 215, Aarhus C 8000, Denmark Hidetaka Furuya E-mail: [email protected] Department of Biological Sciences Graduate School of Science Osaka University Toyonaka, Machikaneyama 1-1, Osaka 560-0043, Japan Kenneth Halanych E-mail: [email protected] Department of Biological Sciences Auburn University 101 Rouse Life Sciences, Auburn, AL 36849, USA Steffen Harzsch E-mail: [email protected] Zoological Institute and Museum–Cytology and Evolutionary Biology University of Greifswald Soldmannstr. 23, House 6.1, Greifswald 17489, Germany Carsten H.G. Müller E-mail: [email protected] Universität Greifswald Zoologisches Institut und Museum Abteilung Allgemeine und Systematische Zoologie Loitzer Str. 26, Greifswald 17487, Germany

Ricardo Neves E-mail: [email protected] Smithsonian Institution National Museum of Natural History 10th Street and Constitution Avenue NW, Washington, DC 20560, USA Yvan Perez E-mail: [email protected] Mediterranean Institute of Marine and Terrestrial Biodiversity and Ecology Aix-Marseille Université (UMR 7263 IMBE) Station Marine d’Endoume Chemin de la Batterie des Lions, Marseille 13007, France Ole Riemann E-mail: [email protected] Rudolf Virchow Center Research Center for Experimental Biomedicine University of Würzburg Josef-Schneider-Str. 2, D15, Würzburg 97080, Germany Andreas Schmidt-Rhaesa (Ed.) E-mail: [email protected] Centre of Natural History Biozentrum Grindel/Zoological Museum Martin-Luther-King-Platz 3, Hamburg 20146, Germany George S. Slyusarev E-mail: [email protected] Department of Invertebrate Zoology Faculty of Biology and Soil Science St. Petersburg State University Universitetskaja nab. 7/9, St. Petersburg 199034, Russia Michael G. Tassia E-mail: [email protected] Department of Biological Sciences Auburn University 101 Rouse Life Sciences, Auburn, AL 36849, USA Oliver Voigt E-mail: [email protected] Department of Earth and Environmental Sciences Ludwig-Maximilians-University of Munich Richard-Wagner-Str. 10, Munich 80333, Germany

Hidetaka Furuya

1 Dicyemida 1.1 Introduction Bodies of animals provide an excellent food and life space for parasites and symbionts. In cephalopods, excretion is carried out by the renal complex (renal and pancreatic appendages) (Fig. 1.1 A) and the branchial heart complex (branchial heart and pericardial appendage). These fluidfilled renal organs, “kidneys”, of cephalopods are unique habitats for the establishment and maintenance of parasites (Hochberg 1982). There are phylogenetically distant parasitic organisms, trematodes, dicyemids, and chromidinids, in the kidney of cephalopods (Nouvel 1945; Hochberg 1983, 1990; Furuya et al. 2004a). Dicyemids (phylum Dicyemida) are the most common parasites of the renal organs of benthic cephalopods, octopuses, and cuttlefishes (Nouvel 1947; McConnaughey 1951; Hochberg 1990; Short 1991; Furuya 1999). Occasionally, dicyemids are found in the pericardium of decapods (Hoffman 1965; Furuya 2007). The body length of dicyemid species ranges from 0.1 to 10 mm. The body of vermiform stages consists of a central cylindrical cell called the axial cell and a single layer of 8 to 40 ciliated external cells called the peripheral cells (Fig. 1.1 C), which are the fewest in number of cells in metazoans except for aberrant myxozoans. The axial cell is a large cell, which extends to 100 mm in length in the largest dicyemid. This organization does not correspond to metazoan two-layered construction of endoderm and ectoderm, and dicyemids have neither body cavities nor differentiated organs. Van Beneden (1877) regarded the dicyemids as intermediate in the body plan between Protozoa and Metazoa and, thus, gave them the name Mesozoa. This phylum included several other microscopical enigmatic organisms, which were not assignable to any other phylum: Trichoplax, Haplozoon, Neresheimeria, Salinella, and orthonectids. Most of these organisms subsequently belonged to the other phyla (Hyman 1949). Only dicyemids and orthonectids were often united into a single phylum Mesozoa. Later Hochberg (1990) and Kozloff (1990) treated them independently as separate phyla, Dicyemida and Orthonectida. However, they were still treated as the Mesozoa in many zoological textbooks because of their unclear relationships to other animals. Several zoologists regard the simple organization of dicyemids to be the result of specialization for parasitism (Nouvel 1947; Stunkard 1954; Ginetsinskaya 1988). However, Hyman (1949), Lapan and Morowitz (1975), and https://doi.org/10.1515/9783110489279-001

Ohama et al. (1984) concurred that dicyemids are primitive multicellular organisms. Because dicyemids have several protozoan-like features, an affinity to the protozoans has been pointed out (Czaker 2006; Noto & Endoh, 2004). Current analyses of molecular sequences have revealed that, rather than truly primitive animals that deserve the name “mesozoan”, they probably belong to the lophotrochozoans (Katayama et al. 1995; Kobayashi et al. 1999; Aruga et al. 2007; Suzuki et al. 2010; Mikhailov et al., 2016). Despite their extremely reduced body plan, dicyemids still appear to exhibit some degree of cell differentiation (Ogino et al. 2011). For this reason, the name Mesozoa is not suited for their phylogenetic place, and Dicyemida, which is the first name of dicyemids introduced by Krohn (1839), has been used as the phylum name since 1999 (Furuya 1999).

1.2 Life cycle The life cycle of dicyemids consists of two phases of different body organization (Fig. 1.2). The first phase includes vermiform stages, in which the dicyemid exists as an asexually formed vermiform embryo, and as a final form, the nematogen or rhombogen. The second phase is the infusoriform embryo that develops from a fertilized egg. A high population density in the cephalopod kidney may cause the shift from an asexual mode to a sexual mode of reproduction (Lapan & Morowitz 1975). Vermiform stages are restricted to the renal sac of cephalopods, whereas the infusoriform embryos escape from the host into the sea to search for a new host. However, it is not clear how infusoriform larvae develop into vermiform stages in the new host. Dicyemids have a high prevalence in their host cephalopods and are usually found to be heavily infecting the renal organs (Fig. 1.1 B). No damage has ever been observed in the infected renal tissue, so dicyemids apparently do no harm to their cephalopod hosts. Lapan (1975a) has even suggested that dicyemids facilitate host excretion of ammonia by contributing to acidification of the urine. In addition to the normal muscular contraction of the renal appendages, the ciliary activity of dicyemids present in the kidneys maintains a constant flow of urine, and as a result dicyemids assist in removal of urine. Thus, dicyemids are symbiotic, rather than parasitic, in their relationship with cephalopods.

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 1 Dicyemida

Fig. 1.1: (A) Schematic diagram of renal organs of octopus. (B) Microphotograph of renal appendages with dicyemids. (C) Whole body of Dicyema japonicum. Bars represent 100 mm in panel B and 50 mm in panel C. AX, axial cell; CL, calotte; D, diapolar cell; G, gill; PA, parapolar cell; RA, renal appendage; RC, renal coelom; RO, renal opening, RS, renal sac; UP, uropolar cell; VC, vena cava.

1.3 Body organization Vermiform stages, vermiform embryos, nematogens, and rhombogens, are similar in shape (Fig. 1.2). The body surface of dicyemids has numerous cilia and a folded structure, which is considered to contribute to absorb nutrients more efficiently from urine (Bresciani & Fenchel 1965; Ridley 1968; Furuya et al. 1997). The number of peripheral cells is species specific and constant. At the anterior region, 4 to 10 peripheral cells form the calotte, of which cilia are shorter and denser than in more posterior peripheral cells (Fig. 1.1 C). The calotte shape varies, depending on the species, and adapts to attach to the various regions of renal tissues in the host kidneys (Furuya et al. 2003a) (Fig. 1.3). Infusoriform embryos are ovoid and have both an anteroposterior and a dorsoventral axis. Embryos mostly consist of 37 or 39 cells (Short 1971; Furuya 1999), which are more differentiated than those of vermiform stages (Matsubara & Dudley 1976; Furuya et al. 2004b). Internally, there are four large cells called urn cells, each containing a germinal cell that probably gives rise to the next generation (Fig. 1.4 C). At the anterior region of the embryo, there is a pair of unique cells called apical cells (Fig. 1.4 E), each containing a refringent body composed of magnesium inositol hexaphosphate (Lapan 1975b). The external cells are mostly ciliated. Infusoriform embryos swim while spinning the body.

The bodies of vermiform stages might be simplified as a reflection of their specialization in their parasitic habitat composed of renal tubules (Nouvel 1947). By contrast, infusoriform embryos seem to represent the true level of organization due to free-swimming organisms (Furuya et al. 1997). However, the body organization of infusoriform embryos cannot be regarded as achieving the grade of tissue level.

1.4 Species diversity of dicyemids To date, 124 dicyemid species have been described from cephalopod hosts distributed in a variety of geographical localities: the Okhotsk Sea, Japan Sea, Western and Eastern North Pacific Ocean, waters around New Zealand, North Indian Ocean, Mediterranean, Western North and Eastern Atlantic Ocean, Gulf of Mexico, and Antarctic Ocean (Catalano 2013; Castellanos-Martinez et al. 2016; Furuya & Tsuneki 2003; Hochberg 1990; McConnaughy 1951; Nouvel 1947; Short 1991). In temperate and polar waters, dicyemids have a very high prevalence (Hochberg 1990; Furuya et al. 2004a). Exceptionally, the prevalence is less than 20% in the cephalopod species that inhabit coral reefs around the tropic and subtropic islands. The phylum Dicyemida includes three families, Conocyemidae, Dicyemidae, and Kantharellae. The number of peripheral cells is species specific and constant in the



1.5 Development of embryos 

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a syncytium (Fig. 1.6 B). In Dicyemidae, the calotte shape is also characteristic of the species of dicyemids (Fig. 1.6 C–G). In the family Dicyemidae, genera are classified by the number of metapolar cells, namely, four cells (Dicyema and Pseudicyema), five cells (Dicyemennea and Dodecadicyema), or six cells (Dicyemodeca) (Fig. 1.5). In the genus Pseudicyema, cells of the propolar tier alternate with cells of the metapolar tier, whereas in the genus Dicyema, the orientation of cells in the calotte is opposite (Fig. 1.5). In the genus Dodecadicyema, two or three micropolar cells present on the top of the propolar tier (Fig. 1.5).

1.5 Development of embryos The development of dicyemids is likely the simplest type of development in the known development of the animal kingdom. Therefore, dicyemids might be useful as a simple model system for studies of cell differentiation and morphogenesis in animals.

1.5.1 Infusoriform embryos

Fig. 1.2: Life cycle of dicyemids. The process involved in the infection of a new cephalopod and the development into the adult are not known (dashed line).

families Conocyemidae and Dicyemidae (Fig. 1.5). The family Kantharellidae contains only one species that is characterized by a variable number of peripheral cells. At the anterior region of dicyemids, 2 to 10 peripheral cells form the calotte. Genera are characterized by the number and orientation of cells in each tier of the calotte (Hochberg 1990). The family Conocyemidae includes two genera, Conocyema and Microcyema; it is characterized by an irregular shape of the adult form. The head of Conocyema polymorpha consists of four cells (calotte) in the embryo (Fig. 1.5), and it looks like a balloon in the adult (Fig. 1.6 A). In Microcyema vespa, the calotte is composed of originally six cells in just formed embryos (Furuya et al. 2001); subsequently, whole peripheral cells are fused into

The infusoriform embryo develops from a fertilized egg. Cell division patterns and cell lineages in the embryogenesis of infusoriform embryos in Dicyema japonicum were known (Furuya et al. 1992a). The fertilized eggs are small, about 12 mm in diameter, and each develops into an infusoriform embryo in the axial cell of rhombogens. In large rhombogens, there can be more than 20 embryos, which include fully formed infusoriform embryos, in the axial cell. The early cleavages are holoblastic and spiral (Fig. 1.7 A–F). The first cleavage is meridional and equal, producing two blastomeres (Fig. 1.7 B). The second cleavage is latitudinal and equal, producing four blastomeres (Fig. 1.7 C). The third cleavage is again equal and results in the eight-cell stage embryo (Fig. 1.7 D). The fourth cleavage is unequal and results in the 16-cell embryo (Fig. 1.7 E). At around the 20- to 24-cell stage, cleavages become asynchronous, and the cleavage pattern changes from spiral to bilateral. The fully formed infusoriform embryo consists of 37 cells and exhibits bilateral symmetry. These 37 cells are produced after only four to eight rounds of cell division. No germ layer is found in the infusoriform embryos of dicyemids, and groups of cells are roughly distinguished as outer and inner cells only. The outer cells on the dorsal and caudal surfaces of the embryo are ciliated.

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 1 Dicyemida

Fig. 1.3: Diagrams of renal organ and dicyemids that have different calotte shapes. B, blood cell; BL, basal lamina; EN, endothelium; ER, epithelium of renal appendage; MC, muscle cell; RC, renal coelom; VC, vena cava.

These cells are derived from the blastomeres of the animal hemisphere of the embryo, and the inner cells are derived from the blastomeres of the vegetal hemisphere. The innermost germinal cells are derived from the cells that form the vegetal pole. Such rearrangements of cells are the basic pattern of the early development of animals. It is also apparent that the outer ciliated cells differentiate much earlier than the inner cells, with the exception of the germinal cells.

cell divides unequally. The anterior large cell becomes an axial cell, whereas the posterior small cell is soon incorporated into the axial cell and becomes an agamete. In D. japonicum, the fully formed vermiform embryo consists of 23 cells and exhibits bilateral symmetry. These 23 cells are produced after only four to six rounds of cell division.

1.5.2 Vermiform embryos

1.6 Gametogenesis in hermaphroditic gonad

Two adult forms of dicyemids, the nematogen and the rhombogen, each develop asexually from an agamete (axoblast) through a larval stage known as the vermiform embryo. The development of vermiform embryos of four genera is known (Furuya et al. 1994, 2001). The early development to the seven-cell stage is conservative, and species-specific differences appear during later stages of embryogenesis (Fig. 1.8). Cell division proceeds spirally in the early stages (Fig. 1.8 A–D). The pattern of cell division beyond the fivecell stage changes from spiral to bilateral (Fig. 1.8 E). After the five-cell stage, divisions occur not one by one but in pairs, and they become almost synchronized. The prospective axial cell is incorporated into the inside of the embryo. At the final stage of embryogenesis, the prospective axial

At the transition stage from the nematogen to the rhombogen, a number of agametes degenerate within the axial cell (Furuya et al. 1993). After an agamete (axoblast) undergoes an unequal first division, excluding a small cell (the paranucleus), the resulting large cell undergoes a second division. Subsequent development depends on the species, and five different types of cell lineage patterns have been recognized (Furuya et al. 1993; Furuya & Tsuneki 2007). In D. japonicum, the larger daughter cell, the progenitor of an infusorigen (Fig. 1.9 A), undergoes an unequal division into two cells. The smaller cell often undergoes an equal division, but one of its daughter cells soon degenerates; the remaining cell is the first spermatogonium (Fig. 1.9 B). The larger cell divides equally to produce the first oogonium and the prospective axial cell of the

1.6 Gametogenesis in hermaphroditic gonad  

 5

Fig. 1.4: Diagrams of the vermiform embryo and the infusoriform embryo. (A, B) Vermiform embryo; (A) cilia omitted; (B) sagittal section; (C–F) infusoriform embryo; (C) sagittal section; (D) dorsal view (cilia omitted); (E) lateral view (cilia omitted); (F) ventral view (cilia omitted). A, apical cell; AG, agamete; AL, anterior lateral cell; AX, axial cell; C, couvercle cell; CA, capsule cell; DC, dorsal caudal cell; DI, dorsal internal cell; E, enveloping cell; G, germinal cell; L, lateral cell; LC, lateral caudal cell; MD, median dorsal cell; MP, metapolar cell; PD, paired dorsal cell; PA, parapolar cell; PP, propolar cell; PVL, posteroventral lateral cell; RB, refringent body; U, urn cell; UP, uropolar cell; VC, ventral caudal cell; VI, ventral internal cell; V1, first ventral cell; V2, second ventral cell; V3, third ventral cell.

infusorigen (Fig. 1.9 C). The axial cell undergoes no further divisions and increases in size. The first spermatogonium is incorporated into the cytoplasm of the axial cell at the three-cell stage (Fig. 1.9 D). The first oogonium, remaining on the periphery of the axial cell of the infusorigen, divides equally to generate a second oogonium and a primary oocyte (Fig. 1.9 E). The first oogonium, remaining at the periphery of the axial cell of the infusorigen, divides equally to generate a second oogonium and a primary oocyte. In the same way, the second oogonium produces a third oogonium and a

primary oocyte (Fig. 1.9 F, G). Thus, oogenesis occurs on the surface of the axial cell of the infusorigen (Fig. 1.9 H). The first spermatogonium within the axial cell of the infusorigen undergoes an equal division to generate a second spermatogonium and a primary spermatocyte (Fig. 1.9 F). In the same way, the second spermatogonium produces a third spermatogonium and a primary spermatocyte. In the axial cell of the maturing infusorigen, two secondary spermatocytes are usually observed in addition to a spermatogonium, a primary spermatocyte and an axial cell nucleus. Soon after the second meiotic division,

6 

 1 Dicyemida

Fig. 1.5: Enface views of the calottes of genera in the two families of Dicyemida (modified from Hochberg 1990). MP, metapolar cell; PR, propolar cell; S, syncytium.

Fig. 1.6: Diversity of dicyemid species. (A) Conocyema polymorpha, (B) Microcyema vespa, (C) Dicyemennea sp., (D) Dicyema ayinense, (E) Dicyema acuticephalum, (F) Dicyemennea sp., (G) Pseudicyema nakaoi.

1.7 Reproductive strategy in the renal organs 

 7

Fig. 1.7: Development of the infusoriform embryo in Dicyema japonicum. (A) A fertilized egg; (B) 2-cell stage; (C) 4-cell stage; (D) 8-cell stage; (E) 16-cell stage; (F) 20-cell stage; (G) 33-cell stage; (H) a formed embryo.

Fig. 1.8: Development of the vermiform embryo in Dicyema japonicum. (A) An agamete; (B) 2-cell stage; (C) 3-cell stage; (D) 4-cell stage; (E) 5-cell stage; (F) 7-cell stage; (G) 9-cell stage; (H) 23-cell stage; (I) a formed embryo.

spermatids are transformed into spermatozoa. Mature spermatozoa are composed of a deeply stained nucleus, and a surrounding small clear area is interpreted as cytoplasm (Fig. 1.9 G, H). The spermatozoon lacks a tail.

1.7 Reproductive strategy in the renal organs Dicyemids most likely evolved from free-living ancestors (Hyman 1940, 1956; Nouvel 1947; Stunkard 1954) with a direct life cycle of several generations in the same host renal organ. The complicated diphasic life cycle of dicyemids, with a characteristic asexual phase, presumably enables them to adapt to their unique habitat in the host renal organs. Asexual reproduction is observed in many endoparasitic groups, including protozoans (Grell 1956; Hochberg 1990; Smyth 1994), cestodes (Hyman 1940, 1949; Stunkard 1975; Rohde 1993), trematodes (Hyman 1940, 1949; Stunkard 1975; Rohde 1993), and orthonectids (Kozloff 1990). Such an asexual reproduction in all the above-mentioned four groups of parasites seems to have

developed independently in each lineage. In these endoparasites, asexual reproduction appears to be an adaptation for similar niches in different hosts. In aquatic animals, taxa with small adults are commonly brooders with embryos held on or in the adult body. However, in species with larger adults, offspring typically are either not cared for or are released at an earlier stage (Strathmann 1990). Adult dicyemids are small in size, and embryos are formed within the adult body. When fully grown, the embryos are released. This essentially equates to brooding. Brooding is common among colonial animals that are composed of many small modules (Strathmann & Strathmann 1982), although brooding style is diverse among bryozoans, pterobranch hemichordates, compound ascidians, and several kinds of hard and soft corals. A population or community of dicyemids formed in the renal sac is similar to a colony, although individuals are monozoic (Furuya et al. 2003b). In dicyemids, the community may develop from a small number of individuals (one or a few) at the initiation of the infection of the renal sac because success of infecting new non-gregarious hosts is apparently low at the level of individual infusoriforms. Dicyemids occasionally are found in only one of the two renal sacs in a host octopus. Two different dicyemid species are occasionally

8 

 1 Dicyemida

Fig. 1.9: Development of infusorigen in Dicyema japonicum. (A) A progenitor cell of infusorigen. (B) Two-cell stage: the small cell is a spermatogonium, the large cell is a mother cell of both oogonium and axial cell of infusorigen. (C) Three-cell stage: nearly two equal-sized cells are the oogonium and axial cell of infusorigen. (D) Three-cell stage: the spermatogonium is incorporated into the axial cell of infusorigen. (E) Four-cell stage: the oogonium divides on the axial cell. (F) Five-cell stage: the spermatogonium is in metaphase. (G) Formed infusorigen. (H) Micrograph of infusorigen: bar represents 10 mm. AI, axial cell of the infusorigen; M, mother cell of the oogonium and axial cell of infusorigen; O, oogonium; PO, primary oocyte; PS, primary spermatocyte; S, spermatogonium; SP, sperm; SS, secondary spermatocyte.

detected, one each in the right and left renal sacs of respective hosts (Furuya et al. 1992b; Furuya 2006). These instances suggest that only a small number of propagules may infect an individual host. Subsequent asexual multiplication forms a large population in the renal sac. Under such conditions, cross fertilization is of little advantage. Thus, dicyemids might undergo self-fertilization via a hermaphroditic gonad. Current studies reveal that dicyemids belong to spiralians. Dicyemids are subject to a number of selecting pressures because of their unique habitat within the kidneys of their cephalopod hosts. In terms of morphological and ecological adaptation, this microenvironment seems to have provided the laboratory space for a simple natural experiment.

Literature Aruga, J., Odaka Y.S., Kamiya A. & Furuya H. (2007): Dicyema Pax6 and Zic: tool-kit genes in a highly simplified bilaterian. BMC Evolutionary Biology 7: 201.

Bresciani, J. & Fenchel T. (1965): Studies on dicyemid Mesozoa. I. The fine structure of the adult (nematogen and rhombogen stage). Videnskabelige Meddelelser Dansk Naturhistorisk Forening 124: 367–408. Castellanos-Martinez, S., Aguirre-Macedo, M. L. & Furuya, H. (2016): Two new species of dicyemid mesozoans (Dicyemida: Dicyemidae) from Octopus maya Voss and Solis-Ramirez (Octopodidae) off Yucatan, Mexico. Systematic Parasitology 93: 551–564. Catalano, S. R. (2013): First descriptions of dicyemid mesozoans (Dicyemida: Dicyemidae) from Australian octopus (Octopodidae) and cuttlefish (Sepiidae) species, including a new record of Dicyemennea in Australian waters. Folia Parasitologica 60: 306–320. Czaker, R. (2006): Serotonin immunoreactivity in a highly enigmatic metazoan phylum, the pre-nervous Dicyemida. Cell and Tissue Research 326: 843–850. Furuya, H. (1999): Fourteen new species of dicyemid mesozoans from six Japanese cephalopods, with comments on host specificity. Species Diversity 4: 257–319. Furuya, H. (2006): Three new species of dicyemid mesozoans (Phylum Dicyemida) from Amphioctopus fangsiao (Mollusca: Cephalopoda), with comments on the occurrence patterns of dicyemids. Zoological Science 23: 105–119.

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Furuya, H. (2007): Redescription of two Dicyemennea (Phylum: Dicyemida) from Rossia pacifica (Mollusca: Cephalopoda: Decapoda). Journal of Parasitology 93: 841–849. Furuya, H. & Tsuneki, K. (2003): Biology of dicyemid mesozoan. Zoological Science 20: 519–532. Furuya, H. & Tsuneki, K. (2007): Developmental patterns of the hermaphroditic gonad in dicyemid mesozoans (phylum Dicyemida). Invertebrate Biology 126: 295–306. Furuya, H., Tsuneki, K. & Koshida, Y. (1992a): Development of the infusoriform embryo of Dicyema japonicum (Mesozoa: Dicyemidae). Biological Bulletin 183: 248–257. Furuya, H., Tsuneki, K. & Koshida, Y. (1992b): Two new species of the genus Dicyema (Mesozoa) from octopuses of Japan with notes on D. misakiense and D. acuticephalum. Zoological Science 9: 423–437. Furuya, H., Tsuneki K. & Koshida, Y. (1993): The development of the hermaphroditic gonad in four species of dicyemid mesozoans. Zoological Science 10: 455–466. Furuya, H., Tsuneki, K. & Koshida, Y. (1994): The development of the vermiform embryos of two mesozoans, Dicyema acuticephalum and Dicyema japonicum. Zoological Science 11: 235–246. Furuya, H., Tsuneki, K. & Koshida, Y. (1997): Fine structure of a dicyemid mesozoan, Dicyema acuticephalum, with special reference to cell junctions. Journal of Morphology 231: 297–305. Furuya, H., Hochberg, F.G. & Tsuneki, K. (2001): Developmental patterns and cell lineages of vermiform embryos in dicyemid mesozoans. Biological Bulletin 201: 405–416. Furuya, H., Hochberg, F.G. & Tsuneki, K. (2003a): Calotte morphology in the phylum Dicyemida: niche separation and convergence. Journal of Zoology 259: 361–373. Furuya, H., Hochberg, F.G. & Tsuneki, K. (2003b): Reproductive traits of dicyemids. Marine Biology 142: 693–706. Furuya, H., Ota M., Kimura R. & Tsuneki K. (2004a): The renal organs of cephalopods: a habitat for dicyemids and chromidinids. Journal of Morphology 262: 629–643. Furuya, H., Hochberg, F.G. & Tsuneki, K. (2004b): Cell number and cellular composition in infusoriform larvae of dicyemid mesozoans (Phylum Dicyemida). Zoological Science 21: 877–889. Ginetsinskaya, T.A. (1988): Trematodes, Their Life Cycles, Biology and Evolution. Amerind Publishing Co. Pvt. Ltd., New Delhi. (Translation of the original Russian edition, 1968) Grell, K.G. (1956): Protozoologie. Springer, Berlin, Göttingen & Heidelberg. Hochberg, F.G. (1982): The “kidneys” of cephalopods: a unique habitat for parasites. Malacologia 23: 121–134. Hochberg, F.G. (1983): The parasites of cephalopods: a review. Memoirs of the National Museum Victoria 44: 109–145. Hochberg, F.G. (1990): Diseases caused by protistans and mesozoans. In: Kinne O. (Ed). Diseases of Marine Animals. Vol. III. Biologische Anstalt Helgoland, Hamburg: 47–202. Hoffman, E.G. (1965): Mesozoa of the sepiolid, Rossia pacifica (Berry). Journal of Parasitology 51: 313–320. Hyman, L. H. (1940): Protozoa through Ctenophora. The Invertebrates. Vol. I. McGraw Hill, New York: 233–247. Hyman, L. H. (1949): Platyhelminthes & Rhynchocoela. The Invertebrates. Vol. II. McGraw Hill, New York: 219–458. Hyman, L. H. (1956): Smaller coelomate groups. The Invertebrates. Vol. V. McGraw Hill, New York: 713–715.

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Katayama, T., Wada H., Furuya H., Sato N. & Yamamoto M. (1995): Phylogenetic position of the dicyemid Mesozoa inferred from 18S rDNA sequences. Biological Bulletin 189: 81–90. Kobayashi, M., Furuya H. & Holland W.H. (1999): Dicyemids are higher animals. Nature 401: 762. Kozloff, E.N. (1990): Invertebrates. Sounders College Publishing, Philaderphia. Krohn, A. (1839): Über das Vorkommen von Entozoen und Kristallablagerungen in den schwammigen Venenanhiängen einiger Cephalopoden. Notizen aus den Gebiete der Natur und Heilkunde 11: 213–216. Lapan, E.A. (1975a): Studies on the chemistry of the octopus renal system and an observation on the symbiotic relationship of the dicyemid Mesozoa. Comparative Biochemistry and Physiology 52: 651–657. Lapan, E.A. (1975b): Inositol polyphosphate deposits in the dense bodies of mesozoan dispersal larvae. Experimental Cell Research 83: 143–151. Lapan, E.A. & Morowitz H.J. (1975): The dicyemid Mesozoa as an integrated system for morphogenetic studies. 1. Description, isolation and maintenance. Journal of Experimental Zoology 193: 147–160. Matsubara, J.A. & Dudley, P.L. (1976): Fine structural studies of the dicyemid mesozoan, Dicyemennea californica McConnaughey. II The young vermiform stage and the infusoriform larva. Journal of Parasitology 62: 390–409. McConnaughey, B.H. (1951): The Life Cycle of the Dicyemid Mesozoa. University of California Press, Berkeley and Los Angeles 55: 295–336. Mikhailov, K.V., Slyusarev G.S., Nikitin M.A., Logacheva M.D., Penin A. A., Aleoshin V.V. & Panchin Y.V. (2016): The genome of Intoshia linei affirms orthonectids as highly simplified spiralians. Current Biology 26: 1–7. Nouvel, H. (1945): Les Dicyémides de quelque Céphalopodes côtes françaises avec indication de la presence de Chromidinides. Bulletin de L’Institut Océanographique, Monaco 887: 1–8. Nouvel, H. (1947): Les Dicyémides. 1re partie: systématique, générations, vermiformes, infusorigène et sexualité. Archives de Biologie, Paris 58: 59–220. Noto, T. & Endoh H. (2004): A “chimera” theory on the origin of dicyemid mesozoans: evolution driven by frequent lateral gene transfer from host to parasite. Biosystems 73: 73–83. Ogino, K., Tsuneki, K. & Furuya H. (2011): Distinction of cell types in Dicyema japonicum (Phylum Dicyemida) by expression patterns of 16 genes. Journal of Parasitology 97: 596–601. Ohama, T., Kumazaki T., Hori T. & Osawa S. (1984): Evolution of multicellular animals as deduced from 5Sribosomal RNA sequences: a possible early emergence of the Mesozoa. Nucleic Acids Research 12: 5101–5108. Ridley, R.K. (1968): Electron microscopic studies on dicyemid mesozoa. I. Vermiform stages. Journal of Parasitology 54: 975–998. Rohde, K. (1993): Ecology of Marine Parasites. CAB International, Wallingford. Short, R. B. (1971): Three new species of Dicyema (Mesozoa: Dicyemidae) from New Zealand. Antarctic Research Series Vol. 17 (Biology of the Antarctic Seas 4): 231–249. Short, R. B. (1991): Dicyemida. Marine Flora and Fauna of the Eastern United States. NOAA Technical Reports NMFS 100.

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Smyth, J.D. (1994): Introduction to Animal Parasitology. Cambridge University Press, Cambridge: 22–154. Strathmann, R.R. (1990): Why life histories evolve differently in the sea. American Zoologists 30: 197–207. Strathmann, R.R. & Strathmann, M.F. (1982): The relationship between adult size and brooding in marine invertebrates. American Naturalists, 119: 91–101. Stunkard, H.W. (1954): The life history and systematic relations of the Mesozoa. Quaternary Review of Biology 29: 230–244.

Stunkard, H.W. (1975): Life-histories and systematics of parasitic flatworms. Systematic Zoology 24: 378–385. Suzuki G.T., Ogino, K., Tsuneki, K. & Furuya H. (2010): Phylogenetic analysis of dicyemid mesozoans (phylum Dicyemida) from innexin amino acid sequences: dicyemids are not related to Platyhelminthes. Journal of Parasitology 96: 614–625. Van Beneden, É. (1876): Recherches sur les Dicyémides, survivants actuels d’un embranchement des Mésozoaires. Bulletins de l’Académie Royale de Belgique 42: 3–111.

George S. Slyusarev

2 Orthonectida 2.1 Introduction Orthonectids are parasites of a wide range of marine invertebrates. They can be found in various taxa of flatworms, nemertines, polychaets, gastropods, clams, ophiurids, and ascidians. Nevertheless, it is a very small group of 25 valid species all in all. The life cycle of orthonectids comprises a parasitic plasmodium occupying the host parenchyma and free-living males and females or hermaphrodites (the latter occurring only in one genus, Stoecharthrum). The males and females develop in the plasmodium and copulate upon their release into the environment. After copulation, the females give birth to larvae, which infect the new host and develop into a new plasmodium. The very first record of orthonectids dates back to 1869 (Keferstein 1869). Five years later, beautiful drawings of orthonectids, although without any description, were published by McIntosh (1874). However, it was Alfred Giard (1877, 1878, 1879, 1880) who first studied and described these enigmatic animals. Metschnikoff (1879a, b, 1881) and Caullery (Caullery 1912, 1914, 1961; Caullery & Lavalee 1908a, b, 1912; Caullery & Mesnil 1899, 1901a, b) also substantially contributed to the studies of this group. Since their first description, orthonectids have been regarded as extremely simple animals with ciliary locomotion, which are devoid of nervous, osmoregulatory, digestive, and muscular systems. Traditionally, orthonectids were believed to lack true tissues and were placed at the same level of organization as sponges (Westheide & Rieger 2007; Brusca & Brusca 2003), i.e., the most primitive Metazoa. However, starting with the studies of Kozloff (1969, 1971, 1990) and thereon (Slyusarev 1994; Hanelt et al. 1996; Pawlowski et al. 1996; Petrov et al. 2010; Mikhailov et al. 2016), it has become more and more clear that orthonectids are not so simple. Female and male orthonectids have been found to possess muscular and nervous systems (Slyusarev & Starunov 2016). It has also been established that orthonectids belong to Bilateria and, more specifically, to highly simplified Spiralia (Mikhailov et al. 2016). Despite obvious advance in our understanding of orthonectids, a number of questions, e.g., the nature of the plasmodium and the way of its formation, still remains open. It is noteworthy that since their discovery, Orthonectida have been joined with Dicyemida in one phylum, Mesozoa (Hartmann 1923; Neresheimer https://doi.org/10.1515/9783110489279-002

1933; Stunkard 1954, 1972; Dodson 1965; Lapan & Morowitz 1972; Remane et al. 1976; Margulis & Schwartz 1998, and many others). However, such attitude casted a lot of doubt, and considering them a separate phylum seems more justified.

2.2 Morphology of free-living females and males 2.2.1 General and external morphology All orthonectids are characterized by exterior sexual dimorphism, manifested in body shape and size (Figs. 2.1 and 2.2 A–C). Females differ from males in body shape and, as a rule, are bigger in size (an exception are males of Rhopalura litoralis, which are of the same size as females but differ in body shape). The body of females is elongated and has a more or less uniform shape, varying from oval to cigarlike (Figs. 2.1, 2.2 A, and 2.3 A, B). The anterior and posterior ends of the body are usually of the same shape (Kozloff 1992). The smallest female (Intoshia variabili) is approximately 75–80 µm long and 15–20 µm across (Alexandrov & Sljusarev 1992), whereas the largest female (Ciliocincta sabellaria) attains 250–265 µm in length and 24–26 µm in width (Kozloff 1965). The longest of all are hermaphrodites of the genus Stoecharthrum, the dimensions of Stoecharthrum monnati being 1625 × 24 µm (Kozloff 1993). Interestingly, two types of females, differing in size and body shape, occur in the genus Rhopalura (Metschnikoff 1881). Female Rhopalura ophiocomae may be either cigarlike, 235–260 µm long and 65–80 µm across, or oval, 125–140 µm long and 65–70 µm across (Fig. 2.1). There is no doubt that both of them belong to the same species. The same phenomenon has been described by Atkins (1933) in Rhopalura granosa. Biological significance of the presence of two forms of females is unclear. Nothing of the kind has been demonstrated in other genera of orthonectids. The body shape of males is much more diverse than that of females (Fig. 2.1). In Intoshia linei, the male is ovoid, the pointed end being the anterior one (Vernet 1990; Haloti & Vernet 1993, 1994); in I. variabili, the body of the male is cylindrical; in R. ophiocomae, the anterior part of the male is noticeably dilated (Fig. 2.1). The size range in

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2.2 Morphology of free-living females and males 

 13

Fig. 2.2: Intoshia variabili. (A) Females, schematic diagrams showing the arrangement of epithelial cells; genital pore (gp). (B) Females, silver impregnation. (C) Male.

males is greater than in females: the tiny male of I. variabili is 35 µm long and 8–9 µm across, whereas the big male of C. sabellaria can reach 125 µm in length and 20–21 µm in width. Both sexes have a genital pore on their ventral side, consisting of five to nine small cells in the midbody. All free-living orthonectids are characterized by the presence of alternating rings of ciliated and non-ciliated cells on their surface (Figs. 2.1 and 2.2 A–C). In most species, the number of the epithelial cells is stable. The highest cell number (620) was registered in the female R. granosa (Atkins 1933), and the lowest one (184) is found in the female I. variabili (Alexandrov & Sljusarev 1992). Among males, the smallest number of epithelial cells is typical for I. variabili. The anterior and the posterior ends of the body are covered with the so-called caps comprising only ciliated cells (Figs. 2.1 and 2.2 A–C). These “caps” are present in both sexes; however, they are more expressed in males. In these zones, the cells are arranged irregularly, and the annular pattern, so characteristic of orthonectids, is not prominent. Regularly alternating rings of ciliated

and non-ciliated cells are located in between the “caps”, producing an effect of distinct “segmentation” in living specimens. This feature permits to easily distinguish living males and females of orthonectids from any protist. Rings of ciliated cells are always wider than those of non-ciliated cells (Figs. 2.1 and 2.2 A–C), the latter always being smaller in size and, as a rule, stretched across the longitudinal axis of the body. Usually, each ring is composed of only one row of cells, but sometimes the rings consist of two or, rarely, three cell rows. It should be noted that besides the ciliated and non-ciliated epithelial cells, the body of all free-living orthonectids comprises muscular cells, nervous cells, and gamets. No other systems are present in orthonectids (Slyusarev 1994; Slyusarev & Starunov 2016).

2.2.2 The structure of epithelial cells Both ciliated and non-ciliated epithelial cells are flattened, their apical and basal surfaces being nearly parallel

◂ Fig. 2.1: Male and female orthonectids, distribution of epithelial cells revealed by silver impregnation, genital pore (gp). The numbers stand for the number of the cells in a ring. After different authors.

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Fig. 2.3: Female Intoshia variabili. (A) SEM, scanning electron microscopy. (B) DIC, differential interference contrast microscopy; cc, ciliated cell; o, oocyte; arrow points to the nervous system. From Slyusarev (1994), with kind permission of Wiley Publishers.

to each other (Figs. 2.4 A–B and 2.5). The epithelial cells are columnar only at the anterior and posterior tips of the body. In silver impregnated specimens, the upper outline of these cells is almost rectangular. In electromicrographs, the adjacent surfaces of the epithelial cells form lateral extensions interlocking like pieces of a jigsaw puzzle (Figs. 2.4 A–B, 2.5, and 2.7 A–B). Ciliated and non-ciliated cells form a typical single-layer epithelium. The apical surface of both ciliated and non-ciliated cells bears short microvilli of irregular shape (Figs. 2.5 and 2.6 A) (Kozloff 1969, 1971; Slyusarev 1994). Microvilli are most prominent in I. variabili, some electron-dense material being present at their tips (Fig. 2.6 A). The epithelial cells are covered with a cuticle, penetrated by the cilia of underlying ciliated cells.

2.2.3 The structure of the cuticle in free-swimming (mature) adults The entire body of mature males and females is covered by a cuticle layer (Slyusarev 2000) (Figs. 2.4 A–B, 2.6 A, and 2.7 A–B). The fine structure of cuticle in males and females

is very similar in various species of orthonectids and differs only in its thickness. The cuticle of I. variabili is approximately 0.34 ± 0.05 µm thick and has the same structure in all parts of the body. It consists of four distinct layers (Fig. 2.6 A–E). The outer layer (layer 1) is thin without any conspicuous structure; the second layer (layer 2) is thicker and appears in sections as a row of fine, discrete, electron-dense granules that are connected with the outer layer. The more solid layer (layer 3), situated under the second one, consists of dense fibers, all arranged parallel to the surface of the cuticle. This layer is continuous to the bottom layer (layer 4) but is not distinctly delimited from it. The bottom layer seems to consist of the same fibers as the previous one, but the fibers are arranged in a kind of meshwork parallel to the cell surface; this is easily seen on sections strictly parallel to this layer and to the cell surface (Fig. 2.6 C–D).

2.2.4 Formation of the cuticle In a single mature plasmodium of I. variabili, all stages of development of males and females are conspicuous

2.2 Morphology of free-living females and males 

 15

Fig. 2.4: Female Intoshia variabili. (A, B) Longitudinal sections of two mature females with oocytes. TEM, transmission electron microscopy; cc, ciliated cell; ci, cilia; cu, cuticle; o, oocyte. From Slyusarev (1994), with kind permission of Wiley Publishers.

(Slyusarev & Miller 1998), which permits to reconstruct the development of the cuticle. At the first stages, the apical surface of the embryo’s epithelial cells is smooth (Fig. 2.8 A). Then the first cilia appear (Fig. 2.8 B) with kinetosomes situated immediately under the cell surface and bearing two cross-striated rootlets. During the same period, the cell shape begins to change and extensive smooth endoplasmic reticulum appears in the cytoplasm.

Alongside with the first cilia, short microvilli-like extensions (0.5 µm long and 0.05 µm mm in diameter) lying flat on the cell surface are formed (Fig. 2.8 C–D). The most characteristic feature of this period of development is that the cisternae of smooth endoplasmic reticulum begin to line up parallel to the borders between the cells, with some of the cisternae coming very close to the cell surface where the cuticle is beginning to form. After that, the extensions

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 2 Orthonectida

Fig. 2.5: Intoshia variabili, schematic diagram showing cell topography in female orthonectid. ci, cilia; cc, ciliated cell; cu, cuticle; o, oocyte. From Slyusarev (1994), with kind permission of Wiley Publishers.

Fig. 2.6: Structure of the cuticle in adult female of Intoshia variabili. (A) General view of the cuticle in cross section of the animal; numbers indicate the layers of the cuticle from the surface to the bottom. (B–D) Sections through cuticle layers 2–4: (B) layer 2; (C) layers 3 and 4; (D) layer 4. (E) Section parallel to cell surface through cell extensions. c, cilia; e, extensions. From Slyusarev (2000), with kind permission of Wiley Publishers. ▸ Fig. 2.7: Intoshia variabili. (A) Transverse section in the anterior fourth of a female with oocytes, showing a ring of muscle cells. (B) Transverse section in the middle of another female, showing interlocking extensions of ciliated cells, muscle cells, granular structures, nuclei, and mitochondria within oocytes. (C–E) mc, muscle cells; cc, ciliated cell; cj, cell junction; cu, cuticle; l, lipid granules; mi, mitochondrion; n, nucleus; o, oocyte. From Slyusarev (1994), with kind permission of Wiley Publishers.

2.2 Morphology of free-living females and males 

 17

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Fig. 2.8: The process of cuticle formation by epithelial cells in Intoshia variabili. (A–E) Development in the plasmodium. (F) Free-swimming adult. c, cilia; cu, cuticle; e, extensions; l, lipid granules; m, microvilli; mi, mitochondria; n, nucleolus; nu, nucleus; p, plasmodial cytoplasm; s, smooth endoplasmic reticulum. From Slyusarev (2000), with kind permission of Wiley Publishers.

on the cell surface grow slightly and gradually bristle up, amorphous material appearing between them. Then they obtain their final shape, which is papilliform with a kind of subdistal bulb or swelling and a narrower tip (Fig. 2.8 E). The extensions are situated very regularly at the surface of the cell. All space between them gets filled with amorphous material, which gradually begins to transform into the layers typical of the adult cuticle (Fig. 2.8 F); this process begins at the surface of the cuticle and progresses toward the inner layers. At the same time, the kinetosomes move down into the small pits typical of the adult ciliated cells. The large nucleolus in the nucleus disappears. The lateral parts of the cell form extensions that interlock with the neighboring cells in jigsaw-puzzle-like manner. During this period, the embryos are covered thick with cilia. Very young embryos seem not to be separated from the plasmodial cytoplasm by a membrane. However, at the time when the first cilia appear on the developing embryo, a membrane separating the embryo from the plasmodial cytoplasm becomes conspicuous around the cilia. Later, when the young adult, using its cilia, begins to move toward the

surface of the host, the membrane appears to be destroyed. With the development of the embryo, the cuticle becomes more dense and compact, and all four cuticular layers become distinct. The regular microvilli-like extensions and endoplasmic reticulum gradually disappear. Finally, the fully developed individuals of I. variabili start moving toward the surface of the plasmodium, using the cilia, and then leave the plasmodium and the turbellarian host.

2.2.5 Cilia In the members of the genera Intoshia and Rhopalura, cilia are evenly distributed on the cell surface, whereas in the genera Ciliocincta and Stoecharthrum, only two rows of cilia are present (Kozloff 1992). In I. variabili and I. linei, cilia are arranged in rows along the anteroposterior axis of the body. In the neighboring rows, the cilia are arranged in a chessboard manner (Fig. 2.9 D). The cilia are 6–7 µm long, and the ciliary axoneme has a typical structure (9 + 2) (Slyusarev & Verulashvili 2005).

2.2 Morphology of free-living females and males 

 19

Fig. 2.9: Ciliated cells in Intoshia variabili. (A) Longitudinal section from the middle of the body (lateral surface uppermost), showing the cuticle; cilia with kinetosomes and rootlets. (B) Transverse section from the first third of the body showing cilia, ring of fibrils, cell junctions, fibrillar structures, tubercles, and matrix. (C) Transverse section showing three-layered cuticle and surface tubercles adjacent to basal layer of cuticle. (D) Section of a ciliated cell parallel to the outer surface and at the level of kinetosomes, showing a band of fibrils, rootlets, kinetosomes, and the cell border. (E) Transverse section showing lamellary bodies. (F) Transverse section showing a pit with cilia. cc, ciliated cell; c, cilia; d, cell junction; cu, cuticle; f, fibrils; r, rootlet; nc, non-ciliated cell; mu, muscle. From Slyusarev (1994), with kind permission of Wiley Publishers.

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Ciliary beating in swimming orthonectids is very well coordinated, and the metachronal wave propagating along the body of the orthonectid is visible in moving specimens. Orthonectids can stop ciliary beating and switch to reversed coordinated beating, which causes orthonectid move with its posterior part foremost. Most often, such reversed ciliary beating happens when the orthonectid hits upon an obstacle, very much like Paramecium.

2.2.6 Cytoskeleton of epithelial cells The cytoskeleton of the epithelial cells consists mainly of fibrous bands (Figs. 2.9 B, D and 2.10) (Slyusarev & Kristensen 2003). In the apical part of the ciliated cell, two fibrous bands, an anterior and a posterior one, each

approximately 0.3 μm thick, run parallel to the cell surface (Figs. 2.9 B and 2.10). These fibrous bands are not only attached to the desmosomes between the cells within a ring of ciliated cells (Fig. 2.9 B, D) but also connect them, traversing the cell. Each band is slightly expanded at its end, where it connects to a spot desmosome. Thus, two fibrous rings are formed within every ring of ciliated cells. The same kinds of fibrous rings, but lying slightly deeper in the cytoplasm, are present within the ring of non-ciliated cells (Fig. 2.10).

2.2.7 The cytoplasm of the ciliated cells The cytoplasm of the ciliated cells has a typical set of cell organelles (Kozloff 1969, 1971; Slyusarev 1994; Slyusarev &

Fig. 2.10: Generalized diagram of cross section of female Intoshia variabili through the fibrous band connecting the cells of one ring. For simplicity, part of the cuticle has been removed, and the number of cilia on the ciliated cells has been reduced. cc, ciliated cell; ci, cilia; d, cell junction; cu, cuticle; f, fibrils; n, nucleus; ru, rootlet. Arrow points to the anterior end of the body. From Slyusarev (1994), with kind permission of Wiley Publishers.

2.2 Morphology of free-living females and males 

Kristensen 2003): the endoplasmic reticulum, free ribosomes, and mitochondria with lamellar cristae (Figs. 2.5, 2.7 A–B, and 2.9 F). In I. variabili and I. linei, mitochondria are often dumbbell shaped, which argues for their division. The dictyosomes of the Golgi apparatus are rare in the epithelial cells of free-swimming specimens, but they are numerous in specimens, which are ready for release from the host but are still in the plasmodium. The electron-lucid cytoplasm contains numerous small and large lipid granules, electron-dense granules, and lamellar bodies (Fig. 2.5). The cytoplasm also contains glycogen (Nouvel 1935a, b, 1939). Each ciliated cell has a small vesicular nucleus (1–1.5 µm) in its basal part. The nucleus encloses decondensed chromatin and a small nucleolus (Figs. 2.5 and 2.10). It should be noted that non-ciliated cells have much less organelles than the ciliated ones, and their cytoplasm often looks electron lucid and “empty” (Fig. 2.9 C). The basis of the ciliary and non-ciliary cells lies either directly on oocytes or sperm cells or rests on muscular cells (Figs. 2.7 B and 2.9 A). The extracellular matrix is poorly developed and does not form any continuous basal membrane between epithelial cells, germ cells, and muscular cells. It is seen as single small patches between the cells. In orthonectids, the extracellular matrix is composed of uniform electron-dense material without any fibers.

2.2.8 The genital pore in a non-fertilized female The genital pore cells arise in the female of I. variabili rather late in the development, after the surface epithelium has completely formed (Slyusarev 2004). At first, they form a compact group of six to seven rounded cells of medium size, located under the surface of the epithelial cells and characterized by the presence of a relatively large vesicular nucleus and numerous granules in the cytoplasm (Figs. 2.11 A–C, 2.12 A, and 2.13 A). In the course of development, the apical surfaces of the genital pore cells are exposed and form a slight depression on the body (Fig. 2.11 C). At the same time, numerous interlacing microvilli are formed on the apical surface of the genital pore cells (Figs. 2.11 B–D and 2.12 B). The cells become more elongated; their basal parts are widened and are in contact with oocytes. Their cytoplasm contains electron-dense secretory granules, mitochondria, lipid inclusions, and a well-developed endoplasmic reticulum. The nuclei of the genital pore cells are located in the basal parts of the cells. A completely formed genital pore of a female is composed of six to seven small cells compactly arranged like segments of an orange. It is noteworthy that a real opening of the

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pore is missing (at least, in I. variabili) because the genital pore cells are tightly apposed (Figs. 2.11 E and 2.12 C). However, the apical surface of the genital pore cells is not covered by a cuticle; it is, in fact, the only place on the whole body that lacks a cuticle (Figs. 2.11 E and 2.12 C). No specialized cellular contacts of the desmosome type can be found between the genital pore cells themselves or between the genital pore cells and the epithelial cells surrounding them, although such contacts are clearly visible between the epithelial cells. The whole process of genital pore formation is completed before the female I. variabili is released from plasmodium where it has developed.

2.2.9 The genital pore in a fertilized female Copulation in I. variabili takes no longer than 30–40 s. After fertilization (oocyte cleavage serving as an indication of successful fertilization), the structure of genital pore cells is changed. The genital pore cells become flattened (Figs. 2.11 E and 2.12 C). The long microvilli on their surfaces disappear, and only the single short ones are left. The secretory granules are no longer visible, and endoplasmic reticulum disappears as well. The nuclei become pycnotic. Observations of living females show that the genital pore serves as a place for the exit of larvae from the female. The first larva leaving the maternal organism pushes the genital pore cells out without damaging the surrounding epithelial ciliated cells and emerges from the female. Within 10–30 min, all the rest of larvae emerge one by one through this opening, while the female goes on swimming.

2.2.10 Musculature Orthonectids are characterized by ciliary locomotion. At the same time, they have been shown to possess a true muscle system (Slyusarev & Manylov 2001; Slyusarev & Starunov 2016). The muscular system of the female I. linei consists only of longitudinal and circular muscles (Figs. 2.13 A and 2.14 A, C). All muscle cells are located immediately under the ciliated ones. The longitudinal muscles lie outside the circular muscles (Fig. 2.7 A, D). The longitudinal musculature comprises two pairs of symmetrical muscle fibers, which start at the anterior end of the worm and run up to its posterior end (Figs. 2.13 A–C and 2.14 A, C). The dorsal and the ventral muscle fibers are of similar structure and run parallel to each other. In all examined specimens, the dorsal muscle fiber curves inward in the

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Fig. 2.11: Genital pore of Intoshia variabili. (A, B) Formation of a genital pore during embryonic development. (C) Genital pore before copulation. (D) Microvilli of the genital pore. (E) Genital pore after copulation. ep, epithelial cell; g, granules; gp, genital pore; m, microvilli; n, nucleus; o, oocyte. From Slyusarev (2004), with kind permission of Folia Parasitologica.

anterior third of the body. In the middle of the body, the ventral muscle bifurcates, forming an oval structure (Figs. 2.13 C and 2.14 C) at the site of the genital pore. At the anterior and the posterior ends of the body, both the dorsal and the ventral muscles bifurcate. At the anterior end, the dorsal and the ventral muscle fibers, diverging and converging again, form a ring perpendicular to the longitudinal axis of the body. This ring embraces exteriorly the termini of the lateral muscles. At the posterior end of the

worm, the dorsal and the ventral muscles bifurcate, forming processes that converge with the right and the left lateral muscles (Fig. 2.14 C). Thus, at the anterior end of the body, the dorsal and the ventral muscles converge, whereas at the posterior end they merge with the lateral ones, forming an integrated longitudinal muscular system. In cross section, the dorsal, the ventral, and the lateral muscle fibers are flattened; the same is true for the circular ones. All longitudinal muscles are 6–7 µm wide throughout.

2.2 Morphology of free-living females and males 

 23

Fig. 2.12: Schematic diagram of the genital pore structure in Intoshia variabili. (А) Genital pore anlagen in embryo. (B) Genital pore before copulation. (C) Genital pore after copulation. ep, epithelial cell; g, granules; gp, genital pore; m, microvilli; n, nucleus; o, oocyte. From Slyusarev (2004), with kind permission of Folia Parasitologica.

The right and the left lateral muscles have similar structure, like the dorsal and ventral ones. At the anterior end, both fibers bifurcate. Thin termini of the lateral muscles serve for their attachment to the ciliated epithelium. Viewed from one angle, such terminus looks just like a fiber extension, whereas from another angle, it appears tapering and pointed at the tip, suggesting that the fiber terminus tapers only in one plane. The muscle fiber

termini are located in between the ciliated epithelial cells. Lateral longitudinal muscles are attached in two spots at the anterior end of the body. The female of I. linei has approximately 9–11 circular muscles (Figs. 2.13 A–C and 2.14 A, C). The circular muscles are nearly perpendicular to the longitudinal ones, parallel to each other throughout the body, and located at about the same distance from one another. The only exception is

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Fig. 2.13: Muscular and nervous systems in female Intoshia linei. (A–C) TRITC-phalloidin staining of muscular system, CLSM. (A) General view, depth-coded image. (B) Lateral view. (C) Ventral view. (D) Cell nuclei stained with DAPI. The ganglion region is outlined. (E) A sagittal section at the level of dorsal (dm) and ventral (vm) muscles, showing serotonin-positive immunoreactivity (green) in the depression of the dorsal muscle. (F) Serotoninergic nervous system, dorsal view. an, anterior nerve; cm, circular muscles; dm, dorsal muscles; gp, genital pore; gc, ganglion cells; lm, lateral muscles; np, ganglionic nerve plexus; pn, posterior nerves; vm, ventral muscle. From Slyusarev and Starunov (2016), with kind permission of Springer Publishers.

the second anterior circular muscle, inclined to the longitudinal axis of the body (Figs. 2.13 B and 2.14 A). The circular muscles are much thinner than the longitudinal ones and still narrower in the sites where they pass under the longitudinal muscle fibers. The muscle system in the female I. variabili (Slyusarev & Manylov 2001) is arranged according to the same plan, as in the female I. linei. Each longitudinal muscle fiber consists of five to seven extremely elongated, overlapping cells, whereas each circular one consists of only three to four such cells (Slyusarev 2003). The cells located at the tips of the longitudinal fibers are especially extended. In the cross sections made in the midbody region, the longitudinal fiber is seen to consist of three to four cells. In the cross sections made at the level of the circular fibers, the longitudinal fibers can be seen outside the muscular circle, inside it, or piercing it through (Figs. 2.9 F and 2.15). The cytoplasm of the muscular cells contains contractile fibrils running along the longitudinal axis of the cell. The fibrils are arranged rather loosely and occupy most of the cell. No cross-striated fibrils have been found. Strangely enough, mitochondria rarely occur in the muscle

cell cytoplasm (Fig. 2.7 E). The nucleus is elongated along the cell axis and is located in the middle part of the cell. True desmosomes have not been observed between the muscular cells or between the muscular and the ciliated cells. The attachment of the longitudinal muscular fibers is rather peculiar (Slyusarev 2003). The terminal cell located at the tip of the muscle fiber is extremely elongated. This cell pierces two or three ciliated cells, the very tip of it deviating toward the surface of the last ciliated cell, where it comes to an end at the level of the cilia rootlets (Fig. 2.5). Entering the cytoplasm of the ciliated cell, the muscular cell invaginates the cell membrane so that the membranes are closely adjacent. No desmosomes can be distinguished between the two membranes. At the very extremity of the muscular cell, there is a thin layer of electron-dense material under the membrane. The ciliated cell cytoplasm surrounding the muscle cell tip is characterized by the presence of loose electron-dense material. No special structures between the ciliated cell and the muscular cell membranes were found. The electron-dense layer of the ciliated cell surrounding the muscular cell tip seems to be connected with the kinetosome rootlets.

2.2 Morphology of free-living females and males 

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Fig. 2.14: Intoshia linei, schematic diagrams of muscular and nervous systems. (A) Muscular and nervous systems, lateral view. (B) Serotoninergic nervous system, dorsal view. (C) Muscular system, ventral view. an, anterior nerve; cm, circular muscles; dm, circular dorsal muscle; gc, ganglion cells; gp, genital pore; lm, lateral muscles; pn, posterior nerves; vm, ventral muscle. From Slyusarev and Starunov (2016), with kind permission of Springer Publishers.

Fig. 2.15: Schematic diagram of the attachment of longitudinal muscles in Intoshia variabili. cc, ciliated cell; F, fibers; R, kinetosome rootlets; N, nucleus; o, oocyte. Arrow shows the direction of ciliary swimming. From Slyusarev (2003), with kind permission of Wiley Publishers.

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2.2.11 The nervous system The entire serotoninergic nervous system as visualized with markers against serotonin of female I. linei is composed of three pairs of cells (Slyusarev & Starunov 2016). All nerve cells lie dorsally. They are multipolar and are compactly located in between the ciliated cells and the muscles (Figs. 2.13 E–F and 2.14 A–B). Two cells (the frontal ones) are shifted anteriorly. Each of them sends a process to the anterior end of the body; this process curves downward and runs along the lateral side of the body. The two largest cells are located centrally (the central cells). Each of them has a long process running backward along the longitudinal muscle fiber and nearly reaching the very end of the body (Fig. 2.14 B). Several processes

A

run forward and connect these cells with the others to form a plexus. Two cells are more lateral (the lateral cells). Each of them sends a process backward, which runs along the longitudinal muscles up to the middle of the body. Two processes of these cells go down and converge to form a ring perpendicular to the longitudinal axis of the body (Figs. 2.15 and 2.16). These cells also have several processes, which run forward and connect them with the others forming the plexus. Thus, the plexus is formed by short processes of all nerve cells and is located between them. The plexus sends a thin unpaired process toward the anterior end of the body, running dorsally. In several examined specimens, this process had serotonin-positive immunoreactivity. However, it is difficult to determine

B

C g

cc

1µm

n n

n

D

1µm

E

1µm Fig. 2.16: Intoshia variabili. Structure of the receptor. (A) Longitudinal section through the receptor showing the nucleus of the receptor cell, the anterior-most oocyte, and ciliated cells. (B–E) Transverse sections through different levels of receptor from several animals. (B) Cilia emerging from the top of the goblet. (D) Middle part of the goblet showing three cells of bowl with cilia inside. (C) Basal part of goblet showing three cells and kinetosomes of the cilia. (E) The stem of the goblet showing the same three cells. cc, ciliated cell; ci, cilia; g, granules; k, kinetosomes; n, nucleus; o, oocyte. From Slyusarev (1994), with kind permission of Wiley Publishers.

2.2 Morphology of free-living females and males 

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which cell it originates. The total number of all nerve cells is no more than 10–12 (see cell counts).

2.2.12 Ciliated receptor A distinctive receptor, composed of three closely apposed cells, is situated along the central axis of the front part of the body, between extensions of ciliated cells of the outer jacket cells and just above the anterior-most oocyte (Figs. 2.16 A and 2.17) (Slyusarev 1994). Together the cells form a tripartite gobletlike structure (Fig. 2.17). The basal part of each cell, containing the nucleus, is widened and situated directly on the anterior-most oocyte. From this base, a long, thin protrusion extends anteriorly, widening into the goblet-shaped anterior part, from the interior of which extend approximately 15 cilia. The cilia have a typical 9 × 2 + 2 structure, with a rootlet starting at the kinetosome. Cilia emerge from the walls of the goblet at a more or less right angle to the wall, then bend upward toward the mouth of the goblet. The cilia appear to intertwine, and those close to the mouth of the goblet appear to emerge less perpendicularly to the longitudinal axis than more posterior ones. At least some of the cilia appear to extend slightly beyond the mouth of the goblet, but none reach the exterior; the receptor is situated under the very small apical cell and between the adjacent six ciliated cells of the first ciliated ring of the anterior end (Figs. 2.16 A–E and 2.17).

2.2.13 Gametes

Fig. 2.17: Intoshia variabili. Reconstruction of receptor and ciliated cells around the goblet; upper part in midsagittal section, middle part (stem), and three-dimensional lower basal part. Drawn by Beth Beyerholm. b, basal part of receptor cell; cc, ciliated cell; ci, cilium/cilia; cu, cuticle; go, goblet of receptor; k, kinetosomes. From Slyusarev (1994), with kind permission of Wiley Publishers.

Gametes are formed in males and females while they are still in the plasmodium. Free-swimming adults have completely developed gametes and can copulate immediately after release from the plasmodium (and the host). Oocytes occupy all space inside the body of the female (Figs. 2.3 B and 2.4 A–B). They lie in a compact mass and sometimes may be arranged in a chessboard order, which is more typical for females of the genus Intoshia, or in an erratic manner, as in female Rhopalura. Oocytes have a large nucleus. The oocyte number in a female varies from 20 to 24 in I. variabili (the smallest of all known orthonectids) up to several hundred (250–300) in Rhopalura. Oocytes are of the same size (11–13 µm in diameter) in all species investigated. Oocytes are rounded and have a centrally located nucleus, which is large and lobe shaped. Their homogeneous cytoplasm contains numerous mitochondria and lipid granules. The spermatozoon structure has been studied in one species only—R. litoralis (Slyusarev & Ferraguti 2002). The

spermatozoon of R.  litoralis is one of the smallest in the animal kingdom. In orthonectids, the process of spermiogenesis takes place while the males are developing inside the plasmodium. The males that emerge spontaneously from the host are mature and can mate immediately. The testis of R. litoralis, situated in the middle part of the body, contains mature spermatozoa and is surrounded by circular and longitudinal muscles. The head of the mature spermatozoon of R. litoralis is slightly elongate, being 1.25 µm long and 0.6–0.7 µm wide (Figs. 2.18 A, C, I and 2.19), and lacks an acrosome. The nucleus appears elongated in longitudinal sections and nearly circular in transverse sections; it has an electron-lucid area in its center. In the midpiece region of the spermatozoon, a single mitochondrion can be observed. In the head region, two centrioles lie under the nucleus in line along the longitudinal axis of the sperm.

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Fig. 2.18: Spermatozoa of Rhopalura littoralis. (A) Longitudinal section. (B) Midpiece region, longitudinal section. (C) Head region with the rootlet, longitudinal section. (D) Proximal centriole, transverse section. (E) Satellite rays, transverse section. (F) Distal centriole, transverse section. (G) Neck region, transverse section. (H) Flagellum, transverse section. (I) Spermatozoa visualized with differential interference contrast (DIC). d, distal centriole; f, flagellum; h, spermatozoon head; m, mitochondrion; n, nucleus; p, proximal centriole; r, rootlet. From Slyusarev and Ferraguti (2002), with kind permission of Wiley Publishers.

2.3 Plasmodium 

 29

The orthonectid sperm also has some specialized features, such as the absence of an acrosome and the presence of a rootlet.

2.2.14 Cell number counts DAPI staining permits to make a precise count of nuclei (hence, the cells) (Fig. 2.13 D). The total number of nuclei in the female of I. linei is 369 ± 2–3. The body of the worm comprises a single-row epithelium, oocytes, muscle, and nerve cells. The number of the epithelial cells is 280. The number of oocytes was counted in living females with DIC and in DAPI-stained specimens, as their nuclei differ significantly in size from all other cells. The oocyte number is 35. Furthermore, the genital pore of the female comprises nine cells. Thus, the total number of epithelial cells, oocytes, and genital pore cells is 280 + 35 + 9 = 324. The rest of the cells (369 – 324 = 45) account for the cells of muscular and nervous systems. The muscle cells lie under the epithelial ones, which permits to count them. The number of muscle cells is 35; hence, there should be only 10 nerve cells. Although the number of all cell types can vary from specimen to specimen, the variation is within a very small range (±2–3), and it does not affect the order of magnitude. Thus, the maximal number of nerve cells should not exceed 10–12. Fig. 2.19: Schematic diagram of a spermatozoon of Rhopalura littoralis. D, distal centriole; F, flagellum; M, mitochondrion; N, nucleus; NF, nuclear fossa; P, proximal centriole; R, rootlet; S, satellite ray. From Slyusarev and Ferraguti (2002), with kind permission of Wiley Publishers.

The nucleus is separated from the proximal centriole by a nuclear fossa (Fig. 2.18 C). The proximal centriole has a length of 0.32 µm and a width of 0.25 µm in the nuclear fossa region; it appears slightly longer than the distal one. The proximal centriole bears a finely cross-striated rootlet (Figs. 2.18 C and 2.19). The rootlet is oriented toward the top of the spermatozoon and runs very close to the nucleus. The cross sections show the satellite rays of the anchoring apparatus, connected to the proximal centriole. The distal centriole has a conventional triplet structure. The basal cross sections of the axoneme show a 9 + 0 structure, which can also be seen in the longitudinal section. Posterior to this short region, the single flagellum has a typical 9 × 2 + 2 structure (Fig. 2.18 D–H). The fine structure of the spermatozoon of R. litoralis generally resembles that of an ectaquasperm (the term suggested by Jamieson 1999), considered “primitive” and plesiomorphic for metazoans.

2.3 Plasmodium Orthonectid plasmodium parasitizes the host parenchyma (Figs. 2.20 B and 2.21 A–D). Generally speaking, the plasmodium can be characterized as a shapeless cytoplasmic sac containing numerous nuclei, cells, and males and females at different stages of development (Metschnikoff 1879a, b; Caullery & Mesnil 1901a, b; Caullery & Lavallee 1912; Atkins 1933; Lang 1954; Caullery 1961; Kozloff 1994, 1997; Slyusarev & Miller 1998). The plasmodium looks very much like an amoeba with pseudopodia of varying length and diameter, which makes it impossible to estimate its size. As a rule, one host houses several individual plasmodia of one species (up to several dozen in I. linei). In the light microscope, although the males and females are very distinct, the borders of the plasmodium are unclear in the living host and rarely visible, which makes counting them difficult. Parasitizing in the host tissues, the plasmodium forms numerous branching fingerlike extensions of varying

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Fig. 2.20: Plasmodium of Intoshia variabili. (A) Macrorhynchus crocea (Turbellaria) infected with a plasmodium of Intoshia variabili. (B) Plasmodium of Intoshia variabili at higher magnification. G, gregarine.

length and diameter. In I. variabili and I. linei, the plasmodium extensions penetrate all parts of the host’s body, including the nervous system. Some extensions break off the maternal plasmodium and give rise to daughter plasmodia. The plasmodium is covered with two distinct plasma membranes (Fig. 2.22 A–C). The external surface of the plasmodium is smooth in some places, whereas it is covered with microvilli in others (Fig. 2.23).

2.3.1 Plasmodial cytoplasm Single plasmodial nuclei are evenly distributed in the plasmodial cytoplasm (Fig. 2.24 A–B), with a slightly higher density near the reproductive cells and embryos. They are usually rounded (approximately 2–2.5 µm in diameter) with a dense, well-defined nucleolus (Fig. 2.24 B). Dense lipid granules and complex granules composed

of smaller rounded granules of approximately 1–2 µm in diameter are quite frequent. Occasionally, very large multivesiculate bodies containing multilaminate structures are scattered within the cytoplasm of the plasmodial extensions (Fig. 2.23). Small mitochondria containing a dense matrix are abundant. Spherical (approximately 2 µm in diameter), dark-staining bodies occur on the borders of the plasmodium. In many cases, the cytoplasm of the plasmodium contains large populations of small vacuoles (Fig. 2.23). A highly developed tubular network resembling smooth endoplasmic reticulum was found in some regions (Slyusarev & Cherkasov 2008).

2.3.2 Reproductive cells In the plasmodial cytoplasm, reproductive cells with well-defined large nuclei can be easily seen. Also,

2.3 Plasmodium 

 31

Fig. 2.21: Plasmodium of Intoshia linei. (A–B) Plasmodium of Intoshia linei inside a living nemertean Lineus ruber. (C–D) Cross sections through the body of the nemertean L. ruber, arrows point to the plasmodium containing males and females at different stages of development. p, plasmodium.

single-cell stages (approximately 3 µm in diameter), twocell stages, which are slightly larger (Fig. 2.24 A–B), and four-cell to many cell stages are apparent. The presence of single and double cells was determined by the method of serial sectioning. Infrequently, reproductive nuclei are found with multivesiculate bodies. With the exception of a few small mitochondria and dense lipid granules of variable sizes (0.3–0.4 µm), the cytoplasm of these cells is amorphous. Occasionally, a single-cell embryo is seen with two nuclei (Fig. 2.24 A). Within the cytoplasm of developing embryos, odd-looking mitotic figures were observed.

2.3.3 Plasmodial feeding In I. linei, the plasmodium feeds by both phagocytosis and pinocytosis (Fig. 2.25 A–B) (Slyusarev & Cherkasov

2008). The similar way of feeding has been observed in I. variabili. Phagosomes are formed only in the sites devoid of microvilli; conversely, the pinocytotic-like vesicles are produced only in the sites covered with microvilli. The domains of the plasmodium plasma membrane seem to be specialized either for phagocytosis or for pinocytosis. It is noteworthy that in 1881, it was Metschnikoff who first proposed the idea about the ability of the orthonectid plasmodium to phagocytize the surrounding host cells. In some species of orthonectids, such as I. leptoplanae, I. linei, I. variabili, and some others, both males and females can develop in one plasmodium, whereas in Rhopalura philinae (Lang 1954) and R. granosa (Atkins 1933; Caullery & Mesnil 1901a), the plasmodium comprises specimens of only one gender. Caullery and Lavallee (1912) and Caullery (1961) proposed calling plasmodia of the first type hermaphroditic and those of the second type

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Fig. 2.22: (A–C) Surface of the plasmodium of Intoshia linei by transmission electron microscopy. n, nucleus; nu, nucleolus; H, host tissue; P, plasmodium. Arrows point to the double membrane of the plasmodium.

Fig. 2.23: Extensions of the plasmodium of Intoshia linei. mi, microvilli; H, host tissue; P, plasmodium.

2.3 Plasmodium 

 33

Fig. 2.24: (A) Part of the plasmodium containing reproductive cells. (A) Dividing reproductive cells. (B) Two plasmodial nuclei and a single reproductive cell in the plasmodial cytoplasm. H, host tissue; ng, reproductive cell nucleus; np, plasmodial nucleus; nu, nucleolus; p, plasmodium. From Slyusarev and Miller (1998), with kind permission of Wiley Publishers.

monosexual. Neresheimer (1933) suggested using terms monoecious and dioecious; Lang (1954) also sustained the latter terminology. In R. ophiocomae and R. granosa, the plasmodium can produce alternately either males or females (Atkins 1933). This might happen when the host is infected with two different monoecious plasmodia, or, probably, dioecious plasmodia can produce males and females interchangeably.

2.3.4 Egress of males and females from the plasmodium It has long been believed by default (Caullery 1961) that completely developed males and females egress from the plasmodium on their own and, moving through the host tissues, finally emerge from the host. Later it was shown (Slyusarev & Cherkasov 2001) that morphological changes of the plasmodium precede the release of adults into the environment. By the time males and females get

fully developed, the surface of the plasmodium forms outgrowths oriented toward the host epithelium. These outgrowths constitute a complicate system of meandering “passages”. The outgrowths penetrate the host tissues, reach the surface of the body, and pierce through the host epithelium in such a way that a small patch of the plasmodium surface comes in touch with the environment. The diameter of the outgrowths gets enlarged, which permits the adults to move inside them. The adults move using ciliary locomotion, and the periods of active motility interchange with periods of rest. The whole process takes from 2 to 10 min.

2.3.5 Plasmodial reproduction The plasmodium can reproduce by budding off new plasmodia (Caullery 1961; Haloti et al. 1992). However, one should admit that this process has not been studied in detail.

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Fig. 2.25: Plasmodial feeding in Intoshia linei. (A) Plasmodium with a well-developed pinocytotic complex. (B) Formation of a phagosome. Arrows show three membranes in the wall of the forming phagosome. l, lipid granule; H, host tissue; f, phagosomes; m, membrane; mi, microvilli; P, plasmodium cytoplasm; S, pinocytotic complex.

2.3.6 The nature of the orthonectid plasmodium It is noteworthy that there are two points of view concerning the nature of the plasmodium. Starting with Giard (1877, 1878, 1879, 1880), Metschnikoff (1879a, b, 1881), and Caullery and Lavallee (1908a, b, 1912), all researchers studying orthonectids believed that the plasmodium is an independent parasitic organism, capable of growth and, probably, of propagation. It develops from a larva upon its invasion into the host tissues. Such attitude remained unchanged until the studies of Kozloff (1990, 1992, 1994, 1997), who proposed an alternative idea: “I am convinced that its (C. sabellaria) plasmodia consist of modified host tissue” (Kozloff 1997, p. 157). Moreover, Kozloff suggested not to use the term “plasmodium” with regard to orthonectids

but to use the word “matrix” instead. A very important detail to be stressed is that, according to Kozloff, the plasmodium lacks its own nuclei, which he has not found (Kozloff 1992, 1994, 1997). In his opinion, numerous plasmodial nuclei described by the previous authors are merely an artifact caused by inadequate techniques. The presence of a parasite triggers hypertrophy of the host cell(s) and leads to formation of a structure, usually referred to as “plasmodium”. Kozloff believes that it is not a whole larva that invades the host, but only individual generative cells. It is still not clear which of these points of view is correct. However, numerous autologous nuclei of the plasmodium have been found in I. linei (Slyusarev & Miller 1998; Slyusarev et al. 2002). The effects of the plasmodium on the host and its pathogenecity have not been studied in detail. For all data available, see Section 2.6.

2.4 Reproduction and development 

2.4 Reproduction and development 2.4.1 Reproductive biology Upon egress from the host, the adult orthonectids survive in seawater (in laboratory conditions) for approximately 2 or 3 days. Females and males demonstrate peculiar sexual behavior; their swimming trajectory is reminiscent of loops (own unpublished results). When the male gets in touch with the female, it attaches to the female’s posterior end. Apparently, the attachment is accomplished because of the interlocking of cilia. For approximately 0.5–3 min, the female drags the male behind itself, and then it bends and grips the male. At this moment, copulation takes place, after which the female straightens up while the male remains attached to its posterior end. For some time, the female continues dragging the male around, trying to liberate itself by shaking its posterior end until it succeeds.

2.4.2 Embryology The data on embryonic development are rather few and fairly contradictory (Giard 1877, 1878, 1879; Metschnikoff 1881; Julin 1881, 1882; Caullery & Mesnil 1901a, b). There is no consensus on this topic among the researchers. So far, the only thing ascertained is that the cleavage is a variety of the spiral cleavage.

2.4.3 Larva In 1901, Caullery and Mesnil published a paper confirming their previous proposal concerning the presence of a migrating stage in the orthonectid life cycle—a ciliated larva infecting a new host (Caullery & Mesnil 1901a, b). The larvae develop synchronously and egress from the genital pore of the female 18–24 h after fertilization. The larva has an oval or, more often, ovoid shape; its length does not exceed 15 µm; and its width is 5 µm (Caullery & Mesnil 1901a, b; Caullery & Lavallee 1908а, b; Atkins 1933; Haloti & Vernet 1995). The number of the cells constituting the larva is estimated to be several dozen. The larvae are all over covered with small ciliated cells with the cilia sometimes longer than the larva itself. Inside the larva, there is a loose mass of several small, supposedly, generative cells. At the anterior-most end of the larva, a group of two to four droplike or pearlike terminal cells is located. These cells are oriented with their pointed ends anteriorly,

 35

and their cytoplasm contains refractile granules. In I. linei, the larva lives no longer than 3 h. In other species, the life span of the larvae is limited to 10–12 h.

2.5 Systematics Systematics of orthonectids cannot be considered completely and finally worked out. The system of orthonectids is based on the distribution of ciliated cells in males and females and on location and number of cilia on the surface of the ciliated cell, revealed by silver impregnation (Nouvel 1935a; Kozloff 1965, 1992). Kozloff (1992) has proposed consecutive numbering of the epithelial cell rings (both ciliated and non-ciliated cells are counted) starting from the anterior end of the body. The number of cell rings varies in different species. In one specimen, the number of cells in different rings may vary. Thus, in R. granosa, the 12th ring comprises 27 cells, whereas in all representatives of the genus Stoecharthrum, the rings consist of only four cells. Phylum Orthonectida Family Rhopaluridae (Stunkard 1937) Genus Ciliocincta (Giard 1877) Ciliocincta akkeshiensis (Tajika 1979)—Hokkaido, Japan; in flatworms (Turbellaria) Ciliocincta julini (Caullery &Mesnil 1899)—E North Atlantic, in polychaetes Ciliocincta sabellariae (Kozloff 1965)—San Juan Islands, WA (USA); in polychaete (Neosabellaria cementarium) Genus Intoshia (Giard 1877) Intoshia leptoplanae (Giard 1877)—E North Atlantic, in flatworms (Leptoplana) Intoshia linei (Giard 1877)—E North Atlantic, in nemertines (Lineus) = Rhopalura linei Intoshia major (Shtein 1953)—Arctic Ocean; in gastropods (Lepeta, Natica, Solariella) = Rhopalura major Intoshia metchnikovi (Caullery & Mesnil 1899)—E North Atlantic, in polychaetes and nemertines Intoshia paraphanostomae (Westblad 1942)—E North Atlantic, in flatworms (Acoela) Intoshia variabili (Alexandrov & Sljusarev 1992)—Arctic Ocean, in flatworms (Macrorhynchus crocea) Genus Rhopalura (Giard 1877) Rhopalura elongata (Shtein 1953)—Arctic Ocean, in bivalves (Astarte)

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Rhopalura granosa (Atkins 1933)—E North Atlantic, in bivalves (Pododesmus) Rhopalura intoshi (Metchnikoff 1881)—Mediterranean, in nemertines Rhopalura litoralis (Shtein 1953)—Arctic Ocean, in gastropods (Onoba aculeus) Rhopalura major (Shtein 1953)—Arctic Ocean, in gastropods (Lepeta, Natica, Solariella) Rhopalura murmanica (Shtein 1953)—Arctic Ocean, in gastropods (Rissoa, Columbella) Rhopalura ophiocomae (Giard 1877)—E North Atlantic, in ophiuroids (usually Amphipholis) Rhopalura pelseneeri (Caullery & Mesnil 1901)—E North Atlantic, polychaetes and nemertines Rhopalura philinae (Lang 1954)—E North Atlantic, in gastropods Rhopalura pterocirri (de Saint-Joseph 1896)—E North Atlantic, in polychaetes Genus Stoecharthrum (Kozloff 1993) Stoecharthrum burresoni (Kozloff 1993)—E North Atlantic in ascidians (Ascidia callosa) Stoecharthrum fosterae (Kozloff 1993)—Arctic Ocean, in bivalves (Mytilus trossulus) Stoecharthrum giardi (Caullery & Mesnil 1899)—E North Atlantic, in polychaetes Stoecharthrum monnati (Kozloff 1993)—E North Atlantic, in mollusks Family Pelmatosphaeridae (Stunkard 1937) Pelmatosphaera polycirri (Caullery &Mesnil 1904)—E North Atlantic, in polychaetes and nemertines It should be noted that some newly described orthonectid species (de Saint-Joseph 1896; Meinkoth 1956; Schilke 1970a, b; Tajika 1979; Haloti & Vernet 1996; and some others) have been published without any silver impregnation images, which makes such descriptions useless and invalid. There are also records of findings of new orthonectids lacking a valid description (Mesnil & Caullery 1918; Foster 1982; etc.). According to Kozloff (1992), not less than 20 orthonectid species are still awaiting proper description; it is very likely that this figure is underestimated.

2.6 Distribution, biology, and ecology Orthonectids rarely occur in nature, and they are difficult to find, which is caused by their two peculiarities. First,

orthonectids are characterized by a highly mosaic distribution of their hosts in nature (Shtein 1953; Kozloff 1992). Second, the percentage of the infected hosts is very low. Thus, Haloti and Vernet (1996) point out that only 0.2% of nemerteans were infected with orthonectids in the vicinity of the Biological Station in Roscoff, and slightly more than 3% in Barfleur (Basse-Normandie). Very rarely, the percentage of infection can be high: Køie (1991) registered from 55% to 92.2% of Phline scabra to be infected with Rhopalura sp. in 1988. Distribution of orthonectids in nature is rather enigmatic and can be called mosaic. Rather often, all persistent attempts to find orthonectids in the place, where they have been once registered, fail (Shtein 1953; Foster 1982; Kozloff 1992; own unpublished results). For example, 5 years after Køie had reported a high rate of infection in Øresund, I have not managed to collect a single infected animal in the same region. Annually, from 1987 to 2016, we have been collecting the turbellarian species Macrorhynchus crocea hosting the orthonectid I. variabili in the Chupa inlet of the White Sea (unpublished data). A detailed description of the procedure has been described in Slyusarev (1994). This turbellarian is common and numerous in the overgrowths of filamentous algae. The sampling was made at seven stations situated at a distance from 500 m to 7 km. During this period, more than 15,000 turbellarians have been collected (non-infected worms were always released back in nature). The infected turbellarians were constantly present during all 23 years of observations at one of the seven stations only; in other places, infected turbellarians were found only three times. The rate of infected M. crocea varied from 0.5% to 1%. The reason of such mosaic distribution is still unclear. For most species, the prevalence of invasion fluctuates within 0.2%–1%. The term intensity of infection cannot be applied to orthonectids; one can state only the degree of host injury (Radner 1982). Thus, in specimens of nemertines infected by a mature plasmodium of I. linei producing adults, 30%, 60%, or even 90% of the host body volume has been shown to be compactly affected. Thus, the parasite can invade only a part of the host body, although most of the species take advantage of the whole space available. The question of infection specificity has been very poorly studied (Radner 1982), so it is impossible to draw any conclusions. R. ophiocomae has been shown to parasitize ophiurans Amphipholis squamata, Ophiotrix fragilis, Ophiura albida, and, possibly, Amphiura elegans (Meinkoth 1956; Fontain 1968; Bender 1972). All other species of this genus demonstrate more strict species specificity.

2.7 The life cycle 

All known orthonectids have been found only in the north hemisphere, specifically in the western part of the Arctic Ocean, in the northeast of the Atlantic Ocean, and in the north of the Pacific Ocean. Metschnikoff (1881) was the first to observe the relationships between the orthonectids and their hosts. He noted that in some infected ophiurans, gonads with mature gamets are preserved. This has also been confirmed for R. ophiocomae by Caullery and Mesnil (1901b). According to Atkins (1933), R. granosa also does not cause castration of its host, the clam Heteranomia squamula. Similarly, Lang (1954) indicated that the orthonectid R. philinae parasitizing the gastropod Philina scabra does not lead to its castration. Kinne (1983) registered the preservation of gonads and their ability to produce normal gamets in clams infected with orthonectids. According to Kozloff (1997), the orthonectid C. sabellaria parasitizing the polychaete Neosabellaria cementarium does not affect its egg production, i.e., does not cause any castration. However, the plasmodium invades the nervous system of the host. Thus, Kozloff (1993) mentioned that the orthonectid S. monnati infects the pedal ganglion of the clam Lucinoma borealis, which, in his opinion, should facilitate the egress of the adult orthonectids from the host. Likewise, Cherkasov and Cherkasova (2002) demonstrated that protrusions of the I. linei plasmodium can penetrate all longitudinal nervous fibers of its nemertine host causing their injury. On the grounds of general considerations, the latter should affect the host behavior; however, no signs of changed behavior of the infected nemertines have been observed. Only one case of joint infection by orthonectids and trematodes has been described in the gastropod P. scabra (Køie 1991). Of all infected mollusks, merely 2.8% were manifesting double infection. The orthonectid I. variabili can invade the host turbellarian, which is already infected with a gregarine (Fig. 2.20 B). However, in 2–7 days, all gregarines disappear (own unpublished results). Rarely, the orthonectid R. litoralis infects the host mollusk Onoba aculeus infested with the trematode sporocysts, but in this case as well, with the course of time the trematodes disappear, which is conspicuous in the dissected gastropod (own unpublished results). All these data suggest that the orthonectid plasmodium drives out the competitor parasites.

2.7 The life cycle Upon invading the host, the orthonectid larva develops into a plasmodium. The plasmodium reproduces by budding (agamic proliferation) and gives birth to males

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and females (Slyusarev 2008). All stages of the embryo development take place inside the maternal plasmodium (Fig. 2.26). The adults are formed from the generative cells, which undergo typical cleavage followed by morphogenesis, which is normally characteristic of the zygote. However, in orthonectids, cleavage of generative cells occurs without preceding fertilization. No maturation divisions of the plasmodial generative cells have ever been observed either. By its genetic consequences, this type of reproduction should be considered as agamic (cloning or replicating genetic copies). However, given that generative cells undergo true embryonic development undistinguishable from the embryonic development of fertilized eggs, they may be considered “parthenogenetic eggs” and the process of plasmodial propagation as apomictic (diploid) parthenogenesis. Males and females born by the plasmodium develop without metamorphosis. They egress from the plasmodium into the water and reproduce by normal amphimixis, which includes gamete maturation and syngamy. Although males lack specialized copulative organs, the insemination is accomplished in such a way that, in fact, it can be considered internal fertilization. Thus, in the course of the orthonectid life cycle, agamic reproduction and apomictic parthenogenesis consistently alternate with amphimixis. Animals in the life cycle of which agamic reproduction alternates with amphimixis are well known; their life cycle is defined as metagenesis. These are cnidarians and some cestodes. The invertebrate taxa with alternating apomictic parthenogenesis and amphimixis, such as trematodes, are also common. Their life cycle is qualified as heterogony. All things considered, the life cycle of orthonectids can be regarded as a combination of metagenesis and heterogony and called “metaheterogony”, a new term that we propose specially for this group.

Acknowledgments Thanks to Dr. Kirill Alexandrov’s good graces, I started studying orthonectids in 1987. I will never forget the support and hospitality of my dear colleagues from different countries, Prof. Reinhardt Møbjerg Kristensen, Prof. D. Miller, and Prof. Marco Ferraguti, whose contribution to these studies cannot be overestimated. I am grateful to O. Manylov, A. Cherkassov, and Yu. Verulashvili, who took part in the studies of orthonectids at different times. I am thankful to Dr. M. Makarov for providing accommodation and facilities for fieldwork at the

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Fig. 2.26: Schematic diagram of the orthonectid life cycle. Nucleus of the plasmodium (N) host (H), reproductive cells (GS), maternal plasmodium (P), and daughter plasmodium (PD).

Barentz Sea. I highly appreciate the assistance of the staff of the Marine Biological Station of Saint-Petersburg State University during my annual expeditions to the White Sea. I would like to express my gratitude to S. Bagrov, from whom I have been collecting materials for many years. Last, but not least, I am indebted to Dr. V. Starunov for his permanent aid and tireless efforts in unraveling the puzzle of orthonectids.

Literature Alexandrov, К.E. &Sljusarev G.S. (1992): A new species of orthonectids, Rhopalura variabeli sp.n. (Mesozoa) from the turbellarian Macrorynchus crocea. Parazitologiia 16: 347–351 [in Russian].

Atkins, D. (1933): Rhopalura granosa sp. nov., an orthonectid parasite of a lamellibranch Heteranomia squamula L., with a note on its swimming behaviour. J. Mar. Biol. Assoc. U.K. 19: 233–252. Bender, K. (1972): The orthonectid, Rhopalura ophiocomae (Giard) found in Ophiotrix fragilis (Abilgaard) and Ophiura albida (Forbes) from Norway. Sarsia 49: 29–32. Brusca, R.C. & Brusca, G.J. (2003): Invertebrates. Sinauer, Sunderland. Caullery, M. (1912): Le cycle évolutif des orthonectides. Verh. VIII. Int. Zool. Kongr. Graz: 765–775. Caullery, M. (1914): Rhopalura pelsneeri C. et M., var. vermiculicola var. nov., orthonectide parasite de Tetrastemma vermiculus. Qtfg. Bull. Soc. Zool. 39: 355–361. Caullery, M. (1961): Classe des orthonectides (Orthonectida Giard, 1877). In: Grasse P.-P. (ed.) Traite de Zoologie. Masson & Ci, Paris. Caullery, M. &Lavallée, A. (1908a): La fécondation et le développement des oeuf des orthonectides. I.—Rhopalura ophiocimae. C. R. Acad. Sci. 146: 40–43.

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 2 Orthonectida

of Intoshia linei affirms orthonectids as highly simplified spiralians. Curr. Biol. 26: 1768–1774. Neresheimer, E. (1933): Mesozoa. In: Grimpe, G. & Wagler, E. (eds.) Die Tierwelt der Nord- und Ostsee, Geest & Portig, Leipzig. Nouvel, H. (1935а): Application des techniques d’impregnation argentique a l’etude systematique des orthonectides. Bull. Biol. France-Belgique 69: 503–507. Nouvel, H. (1935b): Les reserves glycogeniques chez les Orthonectides. Etude de leur evolution. C. R. Acad. Sci. 200: 972–973. Nouvel, H. (1939): Nouvelles observations sur la morphologie des orthonectides. Bull. Soc. Zool. Fr. 64: 262–270. Pawlowski, J., Montoya-Burgos, J.-I., Fahrni, J.F., Wuest, J. & Zaninetti, L. (1996): Origin of the Mesozoa inferred from 18S rRNA gene sequences. Mol. Biol. Evol. 13: 1128–1132. Petrov, N.B., Aleshin, V.V., Pegova, A.N., Ofitserov, M.V. & Slyusarev, G.S (2010): New insight into the phylogeny of Mesozoa: evidence from the 18s and 28S rRNA genes. Moscow Univ. Biol. Sci. Bull. 65: 168–170. Radner, D. N. (1982): Orthonectid parasitism effects on the ophiuroid, Amphipholis squamata. In: M. Lawrence, J., (ed.) Echinoderms: Proceedings of the International Conference Tampa Bay, pp. 395–401. Rotterdam, Balkema. Remane, A., Storch, V. & Welsch, U. (1976): Systematische Zoologie. Stämme des Tierreichs. VEB Gustav Fischer Verlag, Jena. de Saint-Joseph Le Baron (1896): Rhopalura pterocirri, n. sp., orthonectide parasite d’un annelide. Bull. Soc. Zool. Fr. 21: 56–59. Schilke, K. (1970a): Kalyptorhynchia (Turbellaria) aus dem Eulittoral der deutschen Nordseeküste. Helgol. Wiss. Meeresunters. 21: 143–265. Schilke, K. (1970b): Zur Morphologie und Phylogenie der Schizorhynchia (Turbellaria: Kalyptorhynchia). Z. Morph. Tiere 67: 118–171. Shtein, G. A. (1953): Ortonektidy roda Rhopalura Giard nekotorykh molliusko v Barentsova moria. Uchenye Zapiski KareloFinskogo Universiteta, Biologicheskie Nauki 5: 171–206 [in Russian]. Slyusarev, G.S. (1994): Fine structure of the female Intoshia variabili (Alexandrov & Sljusarev) (Mesozoa: Orthonectida). Acta Zool. 75: 311–321. Slyusarev, G.S. (2000): Fine structure and development of the cuticle of Intoshia variabili (Orthonectida). Acta Zool. 81: 1–8. Slyusarev, G.S. (2003): The fine structure the muscle system in the female of the orthonectid Intoshia variabili (Orthonectida). Acta Zoologica 84: 107–111.

Slyusarev, G.S. (2004): Fine structure and function of the genital pore of the female of Intoshia variabili (Orthonectida). Folia Parasitologica 51: 287–290. Slyusarev, G.S. (2008): Phylum Orthonectida: morphology, biology, and relationships to other multicellular animals. Biol. Bull. Rev. 69: 403–427 (in Russian). Slyusarev, G.S. & Cherkasov, A.S. (2001): Analysis of possible mechanisms of the emission of the orthonectids from their hosts. Parazitologiia 35: 338–343 (in Russian). Slyusarev, G.S. & Cherkasov A.S. (2008): Structure and supposed feeding mechanisms of the plasmodium of Intoshia linei (Orthonectida). Invertebr. Zool. 5: 47–51. Slyusarev, G.S. & Ferraguti, M. (2002): Sperm structure of Rhopalura litoralis (Orthonectida). Invertebr. Biol. 121: 91–94. Slyusarev, G.S. & Kristensen, R.M. (2003): Fine structure of the ciliated cells and ciliary rootlets of Intoshia variabili (Orthonectida). Zoomorphology 122: 33–39. Slyusarev, G.S. & Manylov, O.G. (2001): General morphology of the muscle system in the female orthonectid, Intoshia variabili (Orthonectida). Cah. Biol. Mar. 42: 239–242. Slyusarev, G.S., Manylov O.G. & Cherkasov, A.S. (2002): Nuclei in the plasmodium of Intoshia variabili (Orthonectida) as revealed by DAPI staining. Parazitologiia 36: 192–194 [in Russian]. Slyusarev, G.S. & Miller, D.M. (1998): Fine structure of the mature plasmodium of Intoshia variabili (Orthonectida), a parasite of the platyhelminth Macrorhynchus crocea. Acta Zool. 79: 319–327. Slyusarev, G.S. & Starunov, V.V. (2016): The structure of the muscular and nervous systems of the female Intoshia linei (Orthonectida). Org. Dev. Evol. 16: 65–71. Slyusarev, G.S. & Verulashvili, Y.T. (2005): Ciliary transition zone of the orthonectid Intoshia variabili. Parazitologiia 39: 166–170 [in Russian]. Stunkard, H.W. (1954): The life history and systematic relations of the Mesozoa. Q. Rev. Biol. 29: 230–244. Stunkard, H.W. (1972): Clarification of taxonomy in the Mesozoa. Syst. Zool. 21: 210–214. Tajika, K.-I. (1979): A new species of the genus Ciliocincta Kozloff, 1965 (Mesozoa, Orthonectida) parasitic in a marine turbellarian from Hokkaido, Japan. J. Fac. Sci. Hokkaido Univ. 21: 383–395. Vernet, G. (1990): Rhopalura linei, orthonectide parasite de l’heteronemerte Lineus ruber. Cah. Biol. Mar. 31: 251–255. Westheide, W. & Rieger, R. (2007): Spezielle Zoologie. Teil 1: Einzeller und Wirbellose Tiere. 2nd edition, Spektrum Akademischer Verlag, Heidelberg.

Oliver Voigt and Michael Eitel

3 Placozoa 3.1 Introduction

Animals belonging to the phylum Placozoa (Grell 1971a, b) are small, benthic, and exclusively marine. Their body depicts a more or less round, flat ciliated disk of variable shape, with a size from below one to a few millimeters in diameter (Fig. 3.1 A), and lacks any internal or external symmetry planes or axes (Manuel 2009). In unfavorable conditions, also thin thread-shaped forms are frequently observed (Fig. 3.1 B). Typical placozoans move over the substratum in an amoeboid, shape-shifting way. Placozoans have a lower side facing the substratum and an upper side facing the water body. Both sides are delimited by single-layered epithelia, which lack basal membranes. The upper and the lower epithelia enclose an interspace of loosely connected and contractile cells, resulting in a three-layered organization of the placozoan body. The simple gross morphology of the animal is mirrored by the presence of only six somatic cell types, of which two were only discovered recently by a combination of microscopy and immunohistology metods (Smith et al. 2014). Long believed to only comprise the species Trichoplax adhaerens (Schulze 18831), the genetic diversity between different placozoan lineages is high, in some cases exceeding that found between families in other non-bilaterian animals (Voigt et  al. 2004, Eitel et  al. 2018). A considerable size variation and reorganization of genes in the mitochondrial genomes of different placozoan lineages further supports the existence of more than one placozoan species (Signorovitch et al. 2007). Also, on the genome level, gene rearrangements are commonplace, and genetic distance analysis of a large set of nuclear proteins strongly supports the existence of several biological species in the Placozoa. Based on these genomic comparisons, a second species in a new genus was described recently (Eitel et al. 2018). Because placozoans can rapidly reproduce asexually, they can be easily cultured in laboratories. Placozoans have a high regenerative capacity. Pieces of the animal

1 As second species, Treproplax reptans (Monticelli 1893) was formally described and supposedly differed from Trichoplax adhaerens by lacking cilia on its upper side (Monticelli 1893). Because placozoans matching the description of Treptoplax reptans were never found again, the species is not accepted today (Syed & Schierwater, 2002). https://doi.org/10.1515/9783110489279-003

can survive and regrow into a complete animal, given that parts of the outer margin are included (Schwartz 1984). Even a total dissociation into single cells can lead to spontaneous reaggregation, cell sorting, and in some cases generation of a complete and viable animal (Schwartz 1984; Eitel, unpublished). Most studies relied on animals kept in laboratory cultures, and little is known about the placozoan biology in their natural habitat.

3.2 Distribution Placozoans were discovered on the wall of a seawater aquarium containing sample material from the Gulf of Trieste (Schulze 1883). The animals have never been directly observed in their natural habitat. Therefore, reports rely on indirect sampling techniques (Fig. 3.2), i.e., collecting by washing-off substrates like rocks, shells, or algae (Maruyama 2004), or by exposing glass slides in the water column for several days or up to few weeks, which are settled by placozoans (Sudzuki 1977, and most following studies). Knowledge about placozoan distribution is probably far from being complete and strongly biased by sampling efforts in different parts of the world. The worldwide distribution of placozoans has been described in previous publications (Pearse & Voigt 2011; Eitel et al. 2013; Miyazawa & Nakano 2018) and is presented in Fig. 3.3, with addition of previously unpublished findings of placozoans in the Maldives (by slide sampling at Vavvaru, Lhaviyani Atoll, in 2014 and 2015 and at Magoodhoo, Faafu Atoll, in 2016; Voigt, unpublished). Placozoans have been found in the littoral of tropical or subtropical regions in the Mediterranean, the Atlantic, the Red Sea, the Indian Ocean, and the Pacific (Fig. 3.1). Only few reports exist for placozoans from colder waters, for example, from the Atlantic coast of Roscoff, France (von der Chevallerie et  al. 2010), from temperate water in Japan (Maruyama 2004; Miyazawa & Nakano 2018), from Monterey Bay in California (Pearse & Voigt 2011), from the Orbetello Lagoon in Italy (Tomasetti et al. 2005), and from Israel (Eitel & Schierwater 2010). In these cases, the sea surface temperature was between 11°C and 17°C during the sampling period. Long exposure of sampling slides in Antarctic waters failed to provide Placozoa (Pearse & Pearse 1991).

42 

 3 Placozoa

Fig. 3.1: A, A typical placozoan in light microscopy (laboratory culture). B, Thread-shaped placozoans develop under unfavorable culture conditions. The complete body of thread-shaped placozoans or parts of it are still flat and attached to the substratum.

Fig. 3.2: A, A placozoan grazing on a green algae mat on a stone collected at Tenerife, Canary Islands, by Jessica Rach (Eitel & Schierwater 2010). B, Slide sampling of placozoans: microscope slides in open plastic containers are exposed hanging in the water column in a coral reef environment (Lhaviyani Atoll Maldives 2015). C, A placozoan found together with other organisms on a slide shown in panel B after 9 days exposure and additional incubated in seawater for 14 days.

Fig. 3.3: Reports of placozoans. Compiled from previous publications (circles: Pearse & Voigt 2011; Eitel et al. 2013; Tecuatl et al. 2017; Miyazawa & Nakano 2018) and own observations (stars). Map generated with SimpleMappr (Shorthouse 2010).

3.3 Biology 

3.3 Biology 3.3.1 Anatomy Each of the three layers of the placozoan body has a specific organization and composition of cell types.

3.3.1.1 Upper epithelium The upper epithelium is made from flat monociliated cells  with a cell body with the nucleus, which is suspended into the fluid-filled interspace (Fig. 3.4 A1). In between these cells, scattered spherical shiny spheres occur (Fig. 3.4 A2), which are lipid-containing structures of uncertain origin (Grell & Ruthmann 1991). According to some authors, they may be absent at least temporarily (Smith et al. 2014).

3.3.1.2 Lower epithelium The lower epithelium mainly consists of columnar, epidermal so-called cylinder cells (Fig. 3.4 B2), with a single cilium arising from a central ciliary cup. Around the cilium, a “spongy meshwork of fenestrated ledges and folds” (Grell & Benwitz 1981) forms structures that in sections can be mistaken for microvilli and frequently are (erroneously) reported as such (e.g., in Smith et al. 2014). Cylinder cells are by far the most abundant cell type in T.  adhaerens, with roughly 72% of all cells belonging to this cell type (Smith et al. 2014). Larger, non-ciliated lipophil cells (Fig. 3.4 B1) with large spherical inclusions close to the lower surface are occurring dispersed between the cylinder cells. Ciliated gland cells (Fig. 3.4 B3) containing numerous granules are also part of the lower epithelium and occur in higher density close to the margin of the animal (Smith et al. 2014). Gland cells have been originally described to secrete enzymes for extracellular digestion. Recent experiments, however, indicate that they are neurosecretory cells that might control locomotion and feeding behavior (Smith et al. 2014).

3.3.1.3 Intermediate layer In the fluid-filled intermediate layer between the upper and the lower epithelium, two cell types occur: the branching fiber cells (Fig. 3.4 C1) and the crystal cells (Fig. 3.4 C3). Fiber cells possess six or more main processes, which are further

 43

branched. Their extensions connect with the cell  bodies of lower and upper epithelial cells (arrow, Fig.  3.4 C1) or the distal extensions of other fiber cells (Fig. 3.4 C2). Fiber cells potentially form a syncytium, although observations of syncytial connections are sometimes rare in the contact between fiber cells (Grell & Benwitz 1974b; Smith et al. 2014). In the fiber cells, all mitochondria and vesicles of unknown function are forming the mitochondrial complex (Fig. 3.4 C1) (Grell & Ruthmann 1991). The function of these clusters is unknown, but it is reasonable to assume that the clustering of mitochondria supports the contractility of fiber cells. Potentially endosymbiotic bacteria occur in their endoplasmic reticulum of the fiber cells (Grell & Ruthmann 1991). Genetic analyses of sequences from the T. adhaerens genome project revealed that these bacteria belong to the family Midichloriacaea of the Alphaproteobacteria order Rickettsiales (Driscoll et  al. 2013). Crystal cells are found in the outer margin of the animal in the intermediate layer (Fig. 3.4 C4). Each of these cells produces an intracellular rhomboid-shaped birefringent granule of aragonite (Mayorova et al. 2018), which is flanked by mitochondria (Smith et al. 2014). Crystal cells contain a cup-shaped nucleus associated with the cell membrane. The birefringent granules formed by this cell type can be observed by cross-polarized microscopy (Pearse et al. 1994; Fig. 3.5). Sometimes, placozoans only contain few or even no birefringent granules, possibly due to the growth stage of the animal and/or due to the water chemistry. Experiments with placozoans with and without these biominerals suggest that they play a role in gravity sensing (Mayorova et al. 2018): on a vertical surface, placozoans with birefringent granules can maintain their vertical position, whereas placozoans with only few of these crystals tend to move in the direction of the gravity force. A possible mechanism of gravity sensing was suggested: The opening of the cup-shaped nucleus is predominantly facing the outer edge of the animal and in a horizontal position surround the crystal. If tilted vertically, the crystals and the nucleus change their relative position to each other due to the gravitational force. In the crystal cells facing “down”, the crystals move out of the cup-shaped nucleus and come in contact with the cell membrane. Here, so the suggestion, they may trigger mechanosensory receptors, resulting in a movement response (Mayorova et al. 2018). Functional genetic, gene expression as well as morphological studies indicate the existence of a seventh, as yet unidentified, pluri- or totipotent marginal stem cells at the boundary of the upper and lower epithelium (Jakob et  al. 2004; Martinelli & Spring 2004, 2005; Guidi et al. 2011). This presumed cell type, however, awaits further characterization.

44 

 3 Placozoa

3.3 Biology 

Fig. 3.5: Aragonitic birefringent granules are formed in the crystal cells of the intermediate layer. Granules can frequently be observed near the margin of the animals. The pictured animal is folded up in the middle part.

3.3.2 Movement and feeding Placozoa show two types of movement involving ciliary gliding: Larger individuals typically crawl over the substrate by amoeboid, shape-shifting motions; in particular, smaller individuals glide on the substrate with the help the cilia of the lower epithelium without changing their shape (Grell & Ruthmann 1991). In laboratory cultures, very small individuals (40 °N and 40 °S): circumglobal in the southern hemisphere. P. maxima, P. gazellae, Solidosagitta marri, Serratosagitta tasmanica, and P. elegans.

190 

 7 Chaetognatha

Tab. 7.3: Distribution of neritic and semineritic species of chaetognaths. Species Neritic and semineritic Sagitta bombayensis Sagitta helenae Parasagitta chilensis Parasagitta euneritica Parasagitta friderici Parasagitta peruviana Parasagitta popovicii Parasagitta setosa Parasagitta tenuis Aidanosagitta alvarinoae Aidanosagitta bedfordii Aidanosagitta crassa Aidanosagitta delicate Aidanosagitta demipenna Aidanosagitta erythraea1 Aidanosagitta johorensis Aidanosagitta guileri Aidanosagitta meenakshiae Aidanosagitta nairi Aidanosagitta oceania Aidanosagitta tropica Ferosagitta hispida Ferosagitta galerita* Ferosagitta madhupratapi Ferosagitta siamensis Zonosagitta bedoti Zonosagitta littoralis Zonosagitta pulchra

Mediterranean Sea

Atlantic Ocean

Pacific Ocean

+

+

+

+

+

+ +

+ + + + +

+

+ + + + +

+ +

+ + +

+ +

+ + +

+ +

+

+

Indian Ocean

+ + + + + +

+ + +

*Putative Lessepsian migrant species after Terbiyik et al. (2007). 1Red Sea species.

Tab. 7.4: Distribution of oceanic species of chaetognaths. Species Epipelagic Pterokrohnia Arabica Parasagitta elegans Aidanosagitta regularis Aidanosagitta neglecta* Aidanosagitta septata Flaccisagitta enflata Ferosagitta americana Ferosagitta robusta Ferosagitta ferox Serratosagitta bierii Serratosagitta pacifica Serratosagitta pseudoserratodentata Serratosagitta serratodentata Serratosagitta tasmanica Pterosagitta draco Sagitta bipunctata Mesosagitta minima Krohnitta balagopali Krohnitta mutabii

Mediterranean Sea

Atlantic Ocean

Pacific Ocean

+

+ + + + + + + + + + +

+ +

+

+

+ + + + +

+ + + + + +

+ + + +

Indian Ocean

Antarctic Ocean

+ + + + + + + + + + + + + +

+

7.3 Ecology and distribution patterns 

 191

Tab. 7.4: (continued) Species Krohnitta pacifica Krohnitta subtilis Mesopelagic Pseudosagitta gazellae Pseudosagitta lyra Pseudosagitta scrippsae Solidosagitta marri1 Solidosagitta planctonis Solidosagitta zetesios Parasagitta megalophthalma Decipisagitta decipiens Decipisagitta neodecipiens Decipisagitta sibogae Zonosagitta lucida Flaccisagitta adenensis Flaccisagitta hexaptera Eukrohnia bathypelagica Eukrohnia calliops Eukrohnia fowleri Eukrohnia hamata Eukrohnia kitoui Eukrohnia sinica Eukrohnia minuta Xenokrohnia sorbei Bathypelagic Solidosagitta abyssicola Pseudosagitta maxima Caecosagitta macrocephala Eukrohnia bathyantarctica Eukrohnia flaccicoeca Eukrohnia macroneura Eukrohnia proboscidea Bathyspadella edentate Bathyspadella oxydentata Bathybelos typhlops Heterokrohnia alvinae Heterokrohnia angeli Heterokrohnia bathybia Heterokrohnia biscayensis Heterokrohnia curvichaeta Heterokrohnia davidi Heterokrohnia discoveryi Heterokrohnia fragilis1 Heterokrohnia furnestinae Heterokrohnia heterodonta Heterokrohnia longidentata1 Heterokrohnia mirabilis Heterokrohnia mirabiloides Heterokrohnia murina Heterokrohnia wishnerae Archeterokrohnia longicaudata Archeterokrohnia docrickettsae Archeterokrohnia palpifera Archeterokrohnia rubra

Mediterranean Sea +

+

+ + +

Atlantic Ocean

Pacific Ocean

Indian Ocean

+ +

+ +

+ +

+ +

+ + + + + +

+ +

+

+ + +

+

+ + + + + +

+

+ + + + +

+ + +

+ + + + + + + + + +

+

+

+ + + +

+ +

+

+ + +

+ + +

+ +

+

*Putative Lessepsian migrant species after Kehayias et al. (1999). 1Cold-water species.

+

+ + + +

+ + + + +

+ + +

+ +

+

+ + + + + +

+ + + + + + + +

Antarctic Ocean

+

+ +

+

192 

 7 Chaetognatha

Non-belt-shaped patterns include endemic or neritic species with more restricted and fragmented geographical ranges. According to the global latitudinal zonation given by van der Spoel and Heyman (1983), neritic chaetognaths have usually warm water patterns (Pierrot-Bults & Nair 1991). Even these classifications give a good overview of the distribution of planktonic chaetognaths, none of them is fully satisfying because some species can be classified into different categories because of local hydrological specificities and mixed water masses.

7.3.1.2 Benthic species All strictly benthic chaetognaths belong to the Spadellidae (Tab. 7.5). Representatives of Krohnittella (Krohnittellidae)

were sampled close to the sea bottom, and a benthic behavior was suggested for this genus (Bieri 1974). However, because of the scarce records for the two known species of this family, it is difficult to confirm this lifestyle. Benthic Spadellidae have been especially prone to apparent endemism in several areas such as the Caribbean, Canary Islands, Mediterranean Sea, and Antarctic Ocean. However, except for the cosmopolitan S. cephaloptera, insufficient sampling and information are available to speculate on the global distribution of numerous Spadellidae species (Pierrot-Bults & Nair 1991). Moreover, the recent discovery of many new and apparently rare species emphasizes that benthic habitats need to be explored more carefully in order to accurately assess the true species diversity of benthic chaetognaths. Marine caves, interstitial habitats, and deep environments should be given special attention.

Tab. 7.5: Distribution of benthic chaetognath species. Species Epibenthic/neritic Spadella angulata Spadella boucheri Spadella bradshawi Spadella cephaloptera Spadella duverti Spadella gaetanoi Spadella interstitialis Spadella japonica Spadella kappae Spadella lainezi* Spadella ledoyeri* Spadella moretonensis Spadella nunezi Spadella valsalinae Spadella xcalakensis Paraspadella anops* Paraspadella caecafea Paraspadella gotoi Paraspadella johnstoni Paraspadella legazpichessi Paraspadella nana Paraspadella pimukatharos Paraspadella pulchella Paraspadella schizoptera Paraspadella sheardi Mesobenthic Spadella antarctica Spadella birostrata Bathybenthic Spadella equidentata Calispadella alata Hemispadella dauvini Krohnittella boureei Krohnittella tokiokai *Troglobitic species.

Mediterranean Sea

+

Atlantic Ocean

+ +

+

+ +

+

+ +

+ + + +

+ +

+

Pacific Ocean

Indian Ocean

+ + + +

+

+

+ +

+

+ + + +

+ +

+ + + + +

Antarctic Ocean

+

7.3 Ecology and distribution patterns 

7.3.1.3 Endemism and rarely recorded species Endemism of true oceanic species, meaning species confined to a single oceanic province, is rare. Among them, P. gazellae and S. marri are limited to the south of the Antarctic Convergence (David 1965). Eukrohnia calliops was sampled both at meso- and bathypelagic depths and is considered endemic to the Gulf of Mexico (Hernández et al. 2009). Zonosagitta lucida and Flaccisagitta adenensis are restricted to the mesopelagic waters of the Western North Indian Ocean (Casanova 1983). The other oceanic species endemic to the warm Indo-Pacific region are epipelagic. Krohnitta balagopali is only found in the Andaman Sea (Nair et al. 2008; Nair & Gireesh 2010). A. neglecta, Aidanosagitta regularis, and Aidanosagitta septata, as well as F. robusta and F. ferox, are typical of the warm tropical and subtropical waters of the Indo-Pacific (Tokioka 1959; Bieri 1959; Andreú et al. 1989; Pierrot-Bults & Nair 1991). P. scrippsae is found in cold or cool waters throughout the North Pacific Ocean, in the Alaska Gyre, California Current, Bering Sea (Alvariño 1965b; Kotori 1972; 1976), and Kuroshio and Oyashio waters (Johnson & Terazaki 2003). S. bierii has been previously known as endemic to the north western Pacific (Alvariño 1961, 1965b; Fagetti 1968) but is also reported in the intertropical region off the west African coasts (Ducret 1968) and Canary Islands (Hernández-Martin 1991), meaning that the patchy distribution of this oceanic species might be caused by provincialism rather than true endemism. Distribution boundaries of neritic species coincide at the same latitude as the oceanic species. However, there is a higher tendency to endemism for neritic species because of geographically restricted patterns, allopatric isolation, and interplay of local environmental features (PierrotBults & van der Spoel 2003). The Indo-Malayan region and the western parts of the Indian and Pacific Oceans show the greatest diversity of neritic species. Most of the species belonging to Aidanosagitta and Zonosagitta are found in the Indo Pacific tropical and subtropical coastal waters (Tab. 7.3). Sagitta bombayensis is typical of the Indian coastal waters (Nair 1975). The recently described Aidanosagitta meenakshiae is endemic to the Andaman Sea (Nair et al. 2008). More restricted distribution is found for F. galerita observed off Nosy Be (Madagascar) (Dallot 1971), and Ferosagitta madhupratapi (Casanova & Nair 1999) and Aidanosagitta nairi (Casanova & Nair 2002), two species recently found in the Agatti Lagoon (Laccadive Sea). Aidanosagitta erythraea is likely an endemic species of the Red Sea but only one specimen has been caught (Casanova 1985b). There is little endemism described in the eastern part of the Pacific Ocean. Pacific species showing a limited geographical range are concentrated along the

 193

South American region with Parasagitta peruviana and Parasagitta popovicii off the Peruvian coasts (Sund 1961), and Parasagitta chilensis described in Chilean fjords (Villenas & Palma 2006). In the Atlantic basin, P. setosa has a discontinuous geographic distribution over the continental shelf in the Eastern North Atlantic Ocean, Mediterranean Sea, and Black Sea (Peijnenburg et  al. 2006). Sagitta helenae and F. hispida are typical Caribbean neritic species and are found on the edge of the Florida Current (Pierce & Wass 1962; Michel 1984). F. hispida is also frequently recorded from the neritic zone of the central West Atlantic Ocean (Vega-Pérez & Schinke 2011). Numerous cases of benthic species with restricted geographical ranges sometimes associated with endemism have been well documented, for instance, in Spadella ledoyeri from marine caves of the Western North Mediterranean Sea (Casanova 1986d); S. valsalinae, a close, epipsammic relative to cave species found in the photic zone of Northern Adriatic Sea (Winkelmann et  al. 2013); Paraspadella nana, Paraspadella schizoptera, Paraspadella pulchella, and Paraspadella anops, the only cavedwelling species belonging to this genus, from the Caribbean Sea (Michel 1984; Bowman & Bieri 1989); S. nunezi and Spadella lainezi from the Canary Islands (Casanova & Moreau 2004; Casanova et al. 2006); Spadella japonica (Casanova 1993b), Spadella boucheri (Casanova & Perez 2000), and P. gotoi (Casanova 1990) from Japan; the mesopsammic S. interstitialis within Amphioxus sediments in coastal waters of the Italian Islands of Elba and Pianosa in the Ligurian Sea (Kapp & Giere 2005); the mesopsammic S. kappae in Amphioxus sediments in the Channel, North Atlantic, off the french coasts (Roscoff) (Schmidt-Rhaesa & Vieler 2018); Calispadella alata from the Mid-Atlantic ridge (Casanova & Moreau 2005); and Spadella antarctica from Antarctic Ocean (Casanova 1991b). Rarely, species have been recorded from deepwater environment where sampling of planktonic organisms is laborious and often causes damages of the sampled specimens (Terazaki 1991). For instance, single or very few individuals of B. typhlops, K. boureei, Krohnittella tokiokai, and Pterokrohnia arabica were caught in this inevitably destructive way so that in terms of morphology and ecology our knowledge on these species remains fragmentary. In addition, few deep-living species have been the subject of molecular analysis because of lack of specimens properly preserved in alcohol. In particular, the case of the newly discovered, presumably bathybenthoplanktonic species B. oxydentata is of interest because molecular analysis contradicted the previous interpretation exclusively based on morphological data. This bitypic genus is problematic because it exhibits a set of morphological features shared by both Spadellidae

194 

 7 Chaetognatha

and Eukrohniidae (Miyamoto & Nishida 2011). Although Casanova (1999) and Bieri (1991a) placed Bathyspadella within Spadellidae, molecular analysis revealed that B.  oxydentata actually belongs to Heterokrohniidae (Miyamoto & Nishida 2011). Heterokrohniidae species are rarely recorded and are often collected close to the seafloor from deep meso- or more frequently bathypelagic layers, meaning that a number of them adopt a benthoplanktonic lifestyle (Casanova 1986a, 1986b, 1993a). This lifestyle limits the possibility of sampling with conventional plankton nets and rather requires the tricky use of a suprabenthic sledge. Finally, benthic habitats are also relatively unexplored regarding the meso- and meiofauna’s biodiversity. New species and genera are still being described. It is likely that dozens of benthoplanktonic, benthic, and meiobenthic (interstitial) chaetognaths might be discovered in the near future by a broad and appropriate sampling on the surface and within the subsurface layers of the sediments or in the water just above, within highly specific and underexplored habitats such as isolated lagoons and reefs, phytal elements such as seagrass meadows, or phylloids of macroalgae, estuaries, marine caves, hydrothermal vent sites, and deep seafloor.

7.3.2 Diel and ontogenetic vertical migrations Diel and ontogenetic vertical migrations are common among zooplankters and significantly influence their vertical distribution (Pierrot-Bults 1982; Pierrot-Bults & Nair 1991; Vinogradov 1997). Two types of diel vertical migration behavior are known, the normal migration (nocturnal ascent) and the less common reverse migration (nocturnal descent) (Ohman et al. 1983). Light intensity is the main factor controlling diel vertical migration of zooplankton organisms (Forward 1976) and has also been demonstrated to influence upward vertical movement of chaetognaths (Sweatt & Forward 1985a, 1985b). Other environmental factors such as season, lunar and tidal cycles, oxygen, temperature, and salinity as well as biological variables such as food availability, sex, age, and biological rhythms may also affect diel vertical migration of zooplankton (McLaren 1963; Pearre 1974; Ohman et al. 1983; Ohman 1990; Andersen & Nival 1991; Cottier et al. 2006; Harvey et al. 2009; Grigor et al. 2014). So far, a debate persists regarding the adaptive significance of this behavior, but it would be linked to the avoidance of predation and cannibalism, maximization of food intake and fecundity, and strategies associated with horizontal transport (McLaren 1963, 1974;

Longhurst 1976; Ohman et al. 1983; Haney 1988; Ohman 1990; Hill 1994). Several studies have addressed some of these hypotheses in planktonic chaetognaths and reported diurnal patterns in feeding rates associated with diel migration (Nagasawa & Marumo 1972; Pearre 1973; Szyper 1978; Sullivan 1980; Feigenbaum 1982; Øresland 1987). Pearre (1973) determined in P. elegans that diel vertical migrations are related to predation. In the same species, Sullivan (1980) demonstrated a feeding rate significantly higher at night during upward migration. Øresland (1987) observed at different seasons off the coast of Sweden diel feeding regimes in P. setosa and P. elegans with an increase of the predation at night in both species, except during the cold period for P. elegans. However, several authors did not find any day/night differences in feeding rates, for instance, in Parasagitta friderici (Stuart & Verheye 1991), P. setosa (Drits & Utkina 1988), and P. maxima (Sameoto 1987), suggesting that, under certain conditions, these species may feed more or less continuously (Stuart & Verheye 1991). According to Stuart and Verheye (1991), the absence of day/night differences in feeding rates may be correlated with either chaetognaths migrating with their prey or moving from one prey-rich area during the day to another at night. Diel and ontogenetic vertical migrations in oligotrophic environments could be an adaptation to reduce the inter- or intraspecific competition and cannibalism in chaetognaths (Kehayias et al. 1994). This idea was previously mentioned by Pearre (1974), King (1979), and Brodeur and Terazaki (1999) for P. elegans. The diet analysis of epipelagic species in the Eastern Mediterranean Sea showed that a given species of chaetognath either follows prey to an area of high concentration or takes advantage of its migration by waiting for the prey to pass by (Kehayias & Kourouvakalis 2010). Besides food availability and predation avoidance, several works have addressed the effect of hydrographic conditions for delimiting vertical migrations in chaeto­ gnaths. Certain mesopelagic chaetognaths such as D. decipiens and S. planctonis do not reach the surface layers during their vertical diel migrations but may be found at the surface during coastal upwelling events (Bieri 1959; Sund 1964; Fagetti 1968). Conversely, downwelling events may be responsible for the sinking of epipelagic species at great depths in the Florida Current (Stepien 1980; Cheney 1985). According to Duró et  al. (1994) and Duró and Gili (1996), the presence of thermocline and pycnocline helps to explain the short vertical migration patterns of chaeto­ gnaths in the northern Benguela region. Conflicting results have been published on the ability of some chaetognaths to migrate through a thermocline. E. hamata is found mainly below the mixed layer with a vertical distribution restricted

7.3 Ecology and distribution patterns 

below the thermocline (Sullivan 1980). Kotori (1972, 1976) and Terazaki (1992) suggested that a thermocline impedes the upward nocturnal migration of adults of P. elegans in the Bering Sea and in the Kuroshio warm-core ring, respectively. However, Conway and Williams (1988) observed that the thermal tolerance of P. elegans varies with the maturity stages and seasons. By contrast, several authors claimed that this euryhaline species can tolerate a wide range of temperature (Alvariño 1965a) and readily cross the thermocline (Terazaki & Marumo 1979; Sullivan 1980). Downward migration during the day of 19 species is blocked by the oxycline in the Sulu Sea, whereas some of these species, e.g., S. pacifica, F. ferox, A.neglecta, and P. draco, can tolerate a wide range of oxygen levels in the Celebes Sea (Johnson et  al. 2006). Adults of P. setosa in the Black sea can tolerate a wide range of oxygen concentration and temperature during their diel migration (Besiktepe & Unsal 2000). Pronounced and easily detectable vertical diel migrations are usually described in mesopelagic species. Vertical distributions at day and night are well documented for P. elegans. This species performs pronounced vertical diel migrations in many locations and the range of depth is related to the maturity stage, season, and location (e.g., Kotori 1972, 1976; Pearre 1973; King 1979; Sullivan 1980; Terazaki & Miller 1986; Terazaki 1998; Brodeur & Terazaki 1999; Johnson & Terazaki 2003; Harvey et al. 2009). Vertical diel migrations have been mentioned repeatedly, sometimes to a lesser degree, in numerous other chaetognaths in all oceans (Alvariño 1964a, 1965a, 1967b; Pierrot-Bults 1982; Casanova 1999; Nair et al. 2002), but conflicting data have also been reported (see Lyons 1976; Genin et al. 1988; Cheney 1985; Pierrot-Bults & Nair 2010). P. elegans does not dielly migrate in the Arctic environment of Baffin Bay (Sameoto 1987) and in the Gullmar fjord off Sweden (Tönnesson & Tiselius 2005). Lyons (1976) found that chaeto­ gnath species in the central North Pacific exhibit almost no vertical diel migration. Genin et  al. (1988) did not report any significant difference of night-to-day biomass mean ratio off California and considered chaetognaths to be weak migrants. According to Cheney (1985) who studied the chaetognath communities in Western North Atlantic Sea, no vertical diel migrations were detected for K.  pacifica, P. draco, S. bipunctata, S.  helenae, F. enflata, F.  hexaptera, M.  minima, S. serratodentata, E. hamata, K.  subtilis, D. decipiens, P. lyra, P. maxima, S. planctonis, E.  bathypelagica, E. fowleri, and C. macrocephala. In S.  tasmanica, night median depth was exclusively found to be significantly shallower than day median depth only for S. tasmanica. Pierrot-Bults and Nair (2010) did not find any clear indication of vertical diel migration for all the chaetognaths observed in the upper 1000 m of the Western

 195

Sargasso Sea. Detecting vertical diel migration might be problematic because of a lack in resolution caused by restricted and patchy sampling methods, especially at coastal stations or in the case of epipelagic chaetognaths, which remain in a thin layer of shallow waters (Cheney 1985; Bohata & Koppelmann 2013; Al-Aidaroos et al. 2017). Indeed, it has been demonstrated that many epipelagic species could migrate dielly, but only at short distances immediately below the surface (0–50 m), e.g., F. hispida, F.  robusta, K. pacifica, F. enflata, S. bipunctata, S.  bierii, S. serratodentata, S.  tasmanica, P. draco, A. neglecta, P.  setosa, P. friderici, Z. nagae, and M. minima (AlmeidaPrado 1968; Pearre 1974; Pierrot-Bults 1982; Andreú 1992; Gibbons 1992, 1994; Duró & Gili 1996; Terazaki 1996; Besiktep & Unsal 2000; Mutlu 2006; Lie et al. 2012). In order to properly describe vertical diel migrations of small magnitude (10 µm in diameter) and is not present in distal or proximal epidermal cells. However, vacuolated epidermal cells occur in several layers, as is especially evident in the collarette region, and contain an apparatus of tonofilaments encircling the periphery of the cytoplasm. These characters made Ahnelt (1984) suggest that vacuolated epidermal cells may have derived

◂ Fig. 7.15: Cellular organization of the multilayered epidermis of chaetognaths. Original TEM micrographs. A, Semischematic reconstruction of the epidermis from dorsolateral trunk region of Spadella cephaloptera. The epidermis comprises an upper unilayer of distal epidermal cells (dep) and a lower multilayer of flattened proximal epidermal cells (pec). The neuronal plexus integrates a basiepidermal domain (bpaz) with more or less orthogonally arranged neurite bundles (bnb) and an expanded intraepidermal domain (ipaz), including a distal layer of condensed neurites and neuronal somata (ipcs) as well as a network of numerous plexus profiles (ipp) making connection to basal neurite bundles, reproduced from Perez et al. (2014). B–F, Original TEM micrographs. B, D–F, S. cephaloptera; C, Ferosagitta hispida. B, Cross section of spadellid multilayered epidermis showing over 15 layers of proximal epidermal cells from ventrolateral region. C, Sagittid multilayered epidermis in cross section with 2–3 layers of proximal epidermal cells. D, Tangential section of apical region of a distal epidermal cell showing a dense network of vacuoles (v) and secretory granules (sg). E, Aspect of cytosol of distal epidermal cell with abundant Golgi stacks (go). F, Several stacked proximal epidermal cells with intraepidermal pockets filled by condensed fibrillous material (cfm). Further labels: bpp, basiepidermal neuronal plexus profile; tlmu, primary longitudinal (trunk) musculature; ecm, extracellular matrix; nu, nucleus; pez, proximal epidermal zone; rER, rough endoplasmatic reticulum; tfa, tonofilament apparatus.

200 

 7 Chaetognatha

A

2 µm

ecm ipcs nu

ihec

ecm

nu

nu

v

pec

go

bpp

ipp

dec

hmc

sg

B

ecm

mmu

C

pepc

fil

hds pp

pepc C

cu 2 µm

cu

D

E

cu

miv

0,5 µm

sg

rc fc

nu

v

ipp nu

epc

rER

sc

2 µm

ipp

pec

dec hmu ecm

pec

v sg 2 µm

Fig. 7.16: Cellular organization of the integument in certain places of the chaetognath body displaying only one (A–D) or several (2–3) layers of epidermal cells (E–H). A–E, G–H, Original TEM micrographs; F, original LM micrograph. A, Cross section of the hood epidermis being multilayered in its outer portion (to the left) and unilayered in its inner portion (ihec, to the right), the arrows indicate columnar junctions tying together proximal epidermal cells (pec). Interstitial hood musculature (hmc) is framed by the epidermal extracellular matrix (ecm).

7.4 Histological structure, cytology, and functional significance of organ systems 

a

F te

ps

 201

H

ecm

te

lf

tz

lfc

rfi

ecm

G

rc pec

tame

rc tlmu

tlmu

G

ecm

rfi

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20 µm

1 µm

rfi

pec tz

rc

rc rc

tz

rfi

pec rc

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pec

rfi fc rc

fc

pec

rc

rfi 2 µm

◂ Spadella cephaloptera. B, Columnar epidermal cells (pepc) ventral to the mouth of Parasagitta setosa, covered by a thick cuticle (cu). C, Detail of apical region of perioral epidermal cells (pepc) as shown in B (see dashed box) exhibiting microvilli (miv) and hemidesmosomelike connections (hds) to papillar projections (pp) of the internal cuticular face; note the extensive equipment with tonofilaments (fil) attached to papillar projections. P. setosa. D, Incurved glandular epithelium in cross section giving an example for an internalized, functionally highly specialized, and unilayered epidermis in chaetognaths. Clusters of granulated secretory cells (sc) are nested in between agranular epidermal cells (epc). P. setosa. E, Distal aspect of lateral fin of S. cephaloptera in cross section. At either side, distal epidermal cells (dec) overlie the 2–3 layers of proximal epidermal cells (pec). Innermost cell layer is interspersed by ray cells (rc). Black arrow marks an initial stage of merocrine exocytosis of glandular secretion. F, Horizontal histological section of anterior part of the tail. S. cephaloptera. G, Slightly oblique mediohorizontal section of the right lateral fin of S. cephaloptera (section indicated by dashed box in F) showing the elongated profile of the ray cells; note the axial pattern of bundled filaments (rfi) occupying most of the ray cells’ cytoplasm; lower left of the micrograph represents deepest section level by cutting through the fin’s extracellular matrix, called the fin core (fc). H, Detail of the transition zone of the filament apparatus of the ray cells and the epidermal ecm (tz); note the appearance of wedge-shaped mass providing an attachment matrix for paracrystalline filaments. Further labels: a, anus/rectum; bpp, basiepidermal neuronal plexus profile; fc, fin core; go, Golgi stack; hmu, head musculature; ipcs, distal layer of condensed neurites and somata of the intraepidermal neuronal plexus; ipp, intraepidermal neuronal plexus profile; lf, lateral fin; lfc, myoepithelial fells of the lateral field; mmu, mandibular musculature; nu, nucleus; ps, posterior septum; rER, rough endoplasmic reticulum; sg, secretory granule; tame, tail mesentery; te, testes; tlmu, primary longitudinal (tail) musculature; v, vacuole.

202 

 7 Chaetognatha

A

B

sg v nu

ecm tfa

dec

axt

dlmc

syv

axt

mi

ap aec

fil pec

ipcs

C

1 µm

1 µm

D

pec frz

lfc

bl fz

bl

bl lfc ap

ecm

E

fb fz

1 µm

2 µm

bl

fz

mi lfc

ap

fb

1 µm

7.4 Histological structure, cytology, and functional significance of organ systems 

from proximal epidermal cells. The reverse assumption, namely, that flattened proximal epidermal cells derived from vacuolated epidermal cells, appears less likely because, according to those electron microscopic investigations that were conducted on postembryonic, stilldifferentiating chaetognath tissues, the integument of neither Sagittidae nor Spadellidae exclusively comprises vacuolated cells (compare figs. 5 and 15–16 in Shinn & Roberts 1994 and figs. 10.9 and 10.10 in Harzsch et  al. 2015). The function of the collarette remains unclear, but it could participate in buoyancy regulation by increasing the surface/volume ratio (Kapp 1991b, Perez et al. 2001).

7.4.1.4 Further types of epidermal cells In addition to distal and proximal epidermal cells, the chaetognath integument also accommodates receptor cells, neurons, and secretory cells (Figs.  7.18–7.19, 7.24–7.25, and 7.33). All mentioned cell types are usually grouped together in a complex manner forming networks or compact organs. Components of the central and peripheral nervous system are intra- and/or basiepidermal and will be addressed in greater detail in respective chapters below. The same applies to multicellular ciliary sense organs, such as the pair of eyes situated at the dorsoposterior end of the head, the corona ciliata in the neck region and ciliary fence, and tuft organs, which are scattered throughout the epidermis.

7.4.1.5 Oligo- and unilayered epidermis and chitinous structures A cuticle-covered area around the mouth opening forming a ventral cephalic mask presumably protects the epidermis from physical damages when capturing prey. In this area, the chaetognath integument displays columnar epidermal cells (called “tensile cells by Ahnelt

 203

1984) normally equipped with a centered nucleus as well as cisternae of the rough endoplasmic reticulum, Golgi stacks, ribosomes, vacuoles of various sizes, scattered mitochondria, and bundles of tonofilaments extending from apical, peg-like projections of the cuticle down to basal aspect of the cell membrane (Ahnelt 1984; Shinn 1997) (Fig. 7.16 B, C). Other lateral and ventral head localities also show a single layer of columnar epidermal cells devoid of a homogenous cuticle but also containing vertically arranged bundles of tonofilaments (attaching to both apical and basal hemidesmosomes) (Figs.  7.16  D and 7.18 B) and secreting specialized, chitinous extracellular structures such as the (1) lateral and ventral head plates (Figs.  7.3  A–B and 7.18  B), the (2) grasping spines (Figs.  7.18  C and 7.19  A), and (3) the anterior/posterior teeth (Figs. 7.5 A–B and 7.21 A). The head plates are rigid and transparent (e.g., Fig. 7.21 A, B). The electron-lucent plate matrix is made of fibrils and pierced by lines of sometimes filled microcanals. The microcanals are basally invaded by microvilli, which are projected from the columnar epidermal cells in large numbers (Ahnelt 1984; Kapp 1991a; Berezinskaya & Malakhov 1994; Shinn 1997) (Fig. 7.18 B). The mode of secretion forming new plate matrix material is assumed to resemble that found on the head of various lophotrochozoans, such as the chaetae of annelids or the gizzard teeth of bryozoans (Shinn 1997). The lateral head plates are paired and are located dorsolaterally on the chaetognath’s head, covered by the hood. Plate-secreting unilayer of gland cells is continuous with a likewise unilayered epithelium folded in a drop-shaped sac containing numerous granulated epithelial (secretory) cells (Fig. 7.16 D). Lateral head plates are framed anteriorly by a group of the anterior teeth and posteriorly by the fan of grasping spines. The triangular ventral head plate abuts the bases of the grasping spines ventrally. The interconnection to some head muscles let previous authors suggest that the head plates may provide flexibility of the head when cephalic muscles contract and, moreover, may channel and move

◂ Fig. 7.17: Subcellular details of the multilayered epidermis of the dorsal trunk and tail regions in Spadella cephaloptera (A) and Ferosagitta hispida (B–E). Original TEM micrographs. A, Inclusion of a presumably apoptotic epidermal cell (aec) subjacent to unilayer of distal epidermal cells (dec); note the presence of secretory granules (sg) lined up at the apex of the distal epidermal cells. B–E, Ultrastructure of the extracellular matrix (ecm) of the epidermis in cross (B–C) and oblique-tangential (D–E) sections. B, ECM of dorsal trunk region; two axonal terminals (axt) of the basiepidermal neuronal plexus with synaptic vesicles (syv) accumulated toward the basal lamina (bl). C, Thick ECM in dorsolateral tail region. Note the two compartments of the ECM comprising an outer zone devoid of or poor in fibrillous material (frz) as well as an inner zone with extensive arrangement of highly ordered fibrils (fz); tiny peg-like protuberances of the inner basal lamina (ap) serve as anchor joints of cytoskeletal filaments (fil) passing through the lateral field cells (lfc). D, Tangential view of the ECM fibrils. E, Fibrillar zone and basal aspects of lateral field cells in high-power magnification. Note the bundled cytoskeletal filaments (fb) adhering to the peg-like protuberances of the basal lamina. Further labels: dlmc, dorsal longitudinal (primary) muscle cell; ipcs, distal layer of condensed neurites and somata of the intraepidermal neuronal plexus; mi, mitochondrion; nu, nucleus; pec, proximal epidermal cell; tfa, tonofilament apparatus; v, vacuole.

204 

 7 Chaetognatha

A

B bl

axt

1 µm

lpm

cu

miv

ecm dlmc syv pec

epc

bpp

fib

1 µm

cu

C

hmu

ecm ac

D

pec sc

sc pcc ecu

sc

nu

dec v

mell 2 µm

E

dec

oedl psg

sg

axt

syv sg

E F

F mib

sov 2 µm

sg

axt syp

axt 0,5 µm

mib

0,5 µm

7.4 Histological structure, cytology, and functional significance of organ systems 

the grasping spines and teeth (Burfield 1927; John 1933; Kuhl 1938; Hyman 1959; de Beauchamp 1960; reviewed in Shinn 1997; see the Fig. 7.2 C, D in this chapter). Despite their length, diameter, and location on the head, anterior/posterior teeth and grasping spines share a similar internal anatomy, mode of formation, and, most probably, the same chemical components such as α-chitin, zinc, and silicon (Atkins et  al. 1979; Bone et  al. 1983; Ahnelt 1984; Berezinskaya & Malakhov 1994; Shinn 1997). Teeth, however, are completely missing in some species, such as Bathyspadella edentata or representatives of the genus Krohnittella (Shinn 1997). In Eukrohnia species, only the anterior teeth are absent, whereas Krohnitta and few species of the genus Spadella lack the posterior ones. Teeth and grasping spines can be divided into three compartments: the (1) wide base that is secreted by a ring of accessory cells, called anchor cells, and represented by a palisade of electron-lucent columns encompassing the base of the (2) elongated, hollow, either smooth or serrated shaft invaded by distal cytoplasmic projections of the pulp cells clustered below the tooth or spine, and the (3) compact and acute tip of the shaft (smooth or serrated) (Fig. 7.18 C). The cuticle of the tooth or spine shaft exhibits three sublayers. The midlayer is thick, electron-lucent, and framed by a much thinner internal and external sublayer of much higher electron-density (Schmidt 1940; Atkins et  al. 1979; Bone et  al. 1983; Ahnelt 1984; Shinn 1997) (Fig. 7.18 C). During growth, new material is added to the shaft’s proximal inner face. New teeth are added posterolaterally of a given tooth row, whereas new grasping spines are incorporated into the fan from anterior (Kapp 1991a, Shinn 1997). Number and external faces of teeth and grasping spines are variable and are, thus, considered important taxonomic characters (Kapp 1991a). The two fans of zinc-hardened grasping spines are exposed when the hood is retracted preventing the prey from escaping once captured

 205

(Thuesen & Bieri 1987). They are not connected to venom glands directly and have not been observed to pierce the prey to inject venom (Bone et  al. 1983; Thuesen & Bieri 1987; Thuesen 1991), but mouthward-swinging of the grasping spines is used to manipulate and cram the prey into the oral cavity (Thuesen & Bieri 1987; personal observations by C.H.G. Müller & Y. Perez). At least, the grasping spines and teeth have been observed to be capable of penetrating the epidermis (and exoskeleton) of planktonic larvae of various fishes and crustaceans (see fig. 2 in Thuesen & Bieri 1987). Thus, an envenoming of the prey may be initiated and supported by mechanical damage caused (Bieri et al. 1983).

7.4.1.6 Vestibular organs Vestibular-ridge papillae (vestibular organs sensu Kapp 1991a) (Fig.  7.5  A), vestibular pits (Fig.  7.31  C) as well as narial, transvestibular (Fig.  7.19  C–E), and intrabuccal pores also contribute to the structural and functional diversification of the single-layered ventral epidermis of the Chaetognatha (see compilation in Thuesen et  al. 1988a). The shape pattern of the vestibular ridges varies considerably among species (e.g., especially developed in Heterokrohnia spp., see Kapp 1991c), but they all bear papillae that are lined up on the ridges and open in one or several pores (e.g., fig. 3 in Thuesen & Bieri 1987; Thuesen et al. 1988b; Thuesen 1991; Shinn 1997; see also Fig. 7.5 A in this chapter). The ultrastructure of the vestibular-ridge papillae has never been a subject of thorough investigation, but first insights revealed that the single layer of columnar epidermal cells is riddled with putative primary mechano- or chemoreceptor cells, including an elaborate system of microtubules for maintaining cell shape. Furthermore, the epidermal cells project numerous microvilli as well as cilia plugging the pore region (Bone & Pulsford

◂ Fig. 7.18: Examples for cellular and subcellular derivatives of the chaetognath epidermis: extracellular matrix (ecm, A): Ferosagitta hispida; lateral plate (B): Parasagitta setosa; grasping spine (C): P. setosa; ventral gland papillae (D–F): Spadella cephaloptera. Original TEM micrographs. A, Close-up of epidermal ecm and basiepidermal neuronal plexus profiles (bpp) in cross section, including a presumably axonal terminal (axt); note the high abundance of synaptic vesicles (syv) in the transition toward the ecm. B, Cross-cut, single-layered epidermis producing the matrix of the lateral plate (lpm), related epidermal cells are equipped with a dense microvillar brush (miv). C, Cross section of basal region of a grasping spine showing several constitutive epidermal cells filling the pulp cavity (pcc), spine basis is encompassed by an electron-dense cuticular sheath (ecu) secreted by surrounding anchor cells (ac). D, Longitudinal section of a multicellular ventral gland papilla in the transition zone of trunk and tail. Three bottle-shaped secretory cells (sc) are visible surrounded by distal epidermal cells (dec). Apices of secretory cells bear a palisade of columnar secretory granules (psg); note also the presence of cellular inclusions resembling axonal terminals. E–F, Close-ups of the apices of two ventral papillar secretory cells in transverse (E) and longitudinal (F) section, sectors are indicated by dotted lines in D. Presumed synaptic profiles contain vesicles (syv) of the same dimension and electrondensity as seen in A. Extensive bundles of microtubules (mib) pass through cytoplasm. Further labels: bl, basal lamina; cu, cuticle; dlmc, dorsal longitudinal (primary) muscle cell; fib, bundles of tonofilaments; hmu, head musculature; mell, median electron-lucent shaft layer; nu, nucleus; oedl, outer electron-dense shaft layer; pec, proximal epidermal cell; sov, secretion overlay (secreted by dec); v, vacuole.

206 

 7 Chaetognatha

A

mep

glep

br mos

mos

mos es

hco

mes

vsg

lp

mdve

coec mcoa pep

mo

gs

mcop

C

cu

mc

ho

B

50 µm

C

ci

mcoa

dec

src

bb

ecm

nu

tvp

pep

pec

cu

1 µm

D

E

ecm

cu

cu pepc

20 µm

ci

tprc

tvp pepc

E

5 µm

ci

ci 0,5 µm

7.4 Histological structure, cytology, and functional significance of organ systems 

1984). Thuesen et  al. (1988a) detected tetrodotoxin or analogous neurotoxins in the head region of P. elegans and additional species using gas chromatography-mass spectrometry. The venom was assumed to be produced either by chaetognaths directly in a hitherto unknown glandular epithelium (vestibular pit, vestibular papillae, and intrabuccal pores; see Burfield 1927; Bieri et al. 1983; summarized by Thuesen et  al. 1988a) or indirectly, supplied by symbiotic bacteria (presumably Vibrio alginolyticus) the location of which, however, could not be specified (Thuesen & Kogure 1989; Shinn 1997). The vestibularridge papillae were suspected to harbor these bacteria but a clear documentation of tetrodotoxin-producing bacteria, as assumed to be present in the pores, is still missing. Likewise, if present at all, it is doubtful whether these bacteria are symbiotic. The paired vestibular pits can be found anterodorsally of the mouth opening, situated between the most anterior grasping spines and the posterior teeth (fig. 1 in Thuesen et al. 1988b or fig. 2.2. in Kapp 1991a, see also Fig. 7.31 C in this chapter). The epidermis inside the pit is single layered and lacks a cuticle; it contains closely aggregated secretory cells that are filled up almost completely with polymorphic glandular vesicles. These vesicles contain a matrix of grana (fig. 13 in Shinn 1997) and are proposed to release mucous-like substance, which may be (1) adhesive and keep hold of captured prey, (2) lubricating to make it easier to swallow the prey, and/or, but less likely (for details, see paragraph above), may (3) be toxic to envenom the prey (Thuesen et al. 1988a, Shinn 1997). New TEM observations presented in this chapter reveal that the transvestibular pores very likely accommodate ciliated receptor cells (Fig. 7.19 C–E) and are therefore addressed in Section 7.4.2, “Sense Organs”. The pluristratified (multilayered) epidermis is also present but much thinner along the lateral and tail fin(s). Duvert and Salat (1990b) showed the fin rays to consist

 207

of specialized epidermal cells. The elongated ray cells exist in the lower layer of proximal epidermal cells and establish fin rays which extend along the enormously thickened ECM (Figs.  7.16  E, G–H and 7.32  E). The ray cells include a highly ordered horizontal system of tightly adjoined filaments showing two parallel, electron-dense lines and weakly osmiophilic bands in alternating arrangement (Duvert & Salat 1990b; Shinn 1997; and Fig. 7.16 E, G in this chapter). The bundles of filaments are associated with hemidesmosome-like junctions at the basal pole of fin ray cells. Moreover, the ray cell filaments terminate medially at the epidermal ECM to which they are attached by a wedge-shaped anchor matrix (e.g., S. cephaloptera: Fig.  7.16  H). It is assumed that the rigidity of the fins in live animals helps chaetognaths to remain stable in the water by counteracting rotational vectors induced by body motion (Jordan 1992; Shinn 1997). Although devoid of intrinsic musculature, fins can nevertheless be moved, such as by the aid of transverse muscles, which is typical for phragmophoran subtaxa. By adjusting the lateral fin position, directions of locomotion may be regulated (Shinn 1997).

7.4.1.7 Multicellular epidermal glands and solitary secretory cells Multicellular glands and solitary secretory cells are also common. Cephalic ones like the presumably mucoussecreting vestibular pits (Fig.  7.31  C) and intrabuccal pores, or the toxin-releasing vestibular-ridge papillae (Fig. 7.5 A), have already been discussed above. A further secretory organ located ventrally in the neck region is the “ventral gland” of the benthoplanktonic species Xenokrohnia sorbei (discovered by Casanova 1993a). A cytological description of this horseshoe-shaped gland was conducted by Casanova (1993a) and Perez (2000). These

◂ Fig. 7.19: Sense organs including only a single (A) or several (C–E) receptor cells. Original light microscopic (semithin sectioning histology: A, C) and TEM (B, D–E) micrographs. A–C, E, Spadella cephaloptera; D, Spadella valsalinae. Examples from head region. A, Oblique-transverse section of midpart of the head exhibiting cuttings of the mouth opening (mo), sensory domain of the brain (br), lateral plate(s) (lp), right vestibular ganglion (vsg), and grasping spines (gs). B. Close-up of single ciliary receptor from dorsolateral head epidermis. Respective (single) receptor cell (src) is nested within the unilayer of distal epidermal cells (dec), receptive cilium (ci) is protruded from epidermis through deeply invaginated apex of the receptor cell. C, High-power magnified, medioventral aspect of unilayered epidermis next to the mouth (pep), sector indicated exemplarily by dotted rectangle in A; cone-like protuberances of the perioral cuticle (cu) with apical openings represent transvestibular pores (tvp). D, Part of single-layered, perioral epidermis including transvestibular pores (arrows) associated with small clusters of receptor cells (tprc). E, Cilia of clustered receptor cells protrude outside via the transvestibular pore, sector marked as dotted rectangle in D. Further labels: bb, basal (ciliary) body; coec, circumesophageal commissure; ecm, extracellular matrix; es, esophagus; glep, pocketed, glandular epithelium; hco, head coelom; ho, hood (praeputium); mc, main connective; mcoa, constrictor muscle of the mouth (musculus constrictor oris alter); mcop, primary constrictor muscle of the mouth (musculus constrictor oris primus); mdve, external vestibular dilator muscle (musculus dilator vestibuli externus); mep, multilayered cephalic epidermis; mes, musculus expanses superior; mos, mediolongitudinal-oblique head muscles (musculus obliquus superficialis); pep, perioral single-layered epidermis; pepc, perioral epidermal cells.

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studies showed that the glandular epithelium forms two grooves composed of a single type of columnar secretory cell, the cytoplasm of which contains large nuclei surrounded by rough endoplasmic reticulum and many apical electron-dense granules. At the edge of each groove, putative adhesive cells are located that, considering their position, may tightly close the grooves to avoid dilution of the secretions in the seawater (Casanova 1993a). Moreover, serial semithin sections indicated that the ventral gland does not have any connection with the digestive system. The functional significance of this organ is still unknown, but Casanova (1993a) suggested a possible involvement in external digestion. Spadellidae also possess solitary secretory cells or multicellular secretory organs nested in the pluristratified epidermis of the trunk and/or tail region (John 1933; Bone & Pulsford 1978; Ahnelt 1984; Casanova et al. 1995; 2003; Shinn 1997) used for adhesion or adjusting the body position for special functions (Feigenbaum 1976; Casanova 1990). Species of Spadella display numerous, often closely aggregated adhesive papillae (“special attachment cells” in Bone & Pulsford 1978) along the ventral surface of the trunk, starting from the region of the ventral nerve center, and that extend to the posterior tip of the tail where they also scatter over the ventral face of the lateral fins as small knobs (Figs. 7.18 D, 7.35 A, and 7.42 A–B). They are large and abundant in the anterior half of the tail and, in particular, at the transition zone of trunk and tail (e.g., S. cephaloptera: John 1933, Bone & Pulsford 1978, Casanova et  al. 1995; S. valsalinae: Winkelmann et  al. 2013, own observations). The distribution pattern of adhesive papillae coincides with the fact that Spadella spp. usually attach to the substrate with the anterior part of the tail (John 1933). Anatomical data on adhesive papillae are scarce. John (1933) provided a schematic drawing showing several tail papillae of S. cephaloptera in longitudinal section (see his text, fig. 2) documenting highly prismatic secretory cells (“adhesive cells”) with centralized nuclei and

blunt tips. Original TEM data on the same species reveal that the elongated bottle-shaped secretory cells include an extensive system of microtubules in cone-like formation (Fig. 7.18 D). The apex of the secretory cells appears slightly swollen and truncate, and a palisade of highly electron-dense secretory granules is aligned immediately below the apical cell membrane (Fig. 7.18 D–E). Moreover, synaptic terminations of neurites of the neuronal plexus are observed in this apical region (Fig. 7.18 E, F). Generally, profiles of the neuronal plexus are especially abundant at the bottom of the adhesive papillae, and they also reach between the secretory cells. This indicates a direct neuronal control of the secretory activity performed by the adhesive papillae (Thiele & Müller, unpublished observations). Species of the genus Paraspadella have more sophisticated secretory organs, represented by finger- or fringe-shaped, or limb-like appendages (Bowman & Bieri 1989; Casanova et  al. 1995, 2003; Shinn 1997). Their cellular architecture is quite similar to that of the fins, and a pluristratified epidermis is present along with fin rays and a more or less complex “connective tissue”. Based on comparative light and electron microscopic data, Casanova et  al. (1995, 2003) assumed that the structural and functional disparity of these adhesive appendages in Spadellidae may be considered a good example for an evolutionary transformation that took place in this chaetognath subtaxon. Their scenario proposes the numerous, strictly epidermal ventral gland papillae (see previous paragraph) in species of Spadella to be the plesiomorphic state for Spadellidae (e.g., Fig. 7.35 A), whereas the large, stout, and bilobed appendages of P. gotoi, which are extensions of the lateral and tail fins (Casanova 1990) and help to rapidly erect the body into vertical position (Goto & Yoshida 1985), could represent the highest derived state. The six elongated and thin finger-like processes of P. schizoptera may represent the plesiomorphic state of the ventral adhesive organs in this genus. Transformations apply to the reduction and

▸ Fig. 7.20: Position and ultrastructure of the retrocerebral organ in Spadella cephaloptera. A, Parasagittal semithin (LM) section of the head. The retrocerebral organ (ro) is embedded into the posterior (sensory) domain of the brain (br). Original. B–F, Original TEM micrographs. B, Longitudinal (parasagittal) view of the retrocerebral organ showing receptor cell somata (rcso) encircling a central cavity completely filled by strongly branched cilia (bci). C, Close-up of the apex of a receptor cell, presence of a basal body (bb), microtubules (mic), and immediately branching axonema (arrow) demonstrate the ciliary nature of the numerous cytoplasmic inclusions in the cavity. Longitudinal section. D, Horizontal section of the cavity and branched ciliary profiles, complementary to C; two obliquely sectioned axonemas (axo) are visible. E, Longitudinal (parasagittal) sections of receptor cell somata are seen stacked onto each other; note the presence of ciliary ultrastructures like the axonema, basal body, and cross-striated ciliary rootlet (cr); axon bundles (axb) project posteriorly. F, Horizontal section of the retrocerebral duct lumen (rcdl) passing through the dorsal epidermis. Duct contains microvillar processes of the retrocerebral duct cells (rcdc) and enflated, polymorphic, electron-lucent inclusions (inc) of uncommon origin. Further labels: cu, cuticle; ecm, extracellular matrix (of the epidermis); es, esophagus; fshc, strongly flattened sheath cells; hco, cephalic coelomic cavity; mep, multilayered head epidermis; mes, musculus expansus superior; mi, mitochondrion; mos, mediolongitudinal-oblique head muscles (musculus obliquus superficialis); nu, nucleus; pec, proximal epidermal cells; pep, single-layered perioral epidermis.

7.4 Histological structure, cytology, and functional significance of organ systems 

A

B

B

mep

pep

mes

fshc rcso

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br mos

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nu bci

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bb bci

axo mic

mi

mic bci

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rcdc miv

axb nu

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 7 Chaetognatha

A

B

at

lp mos

ho

ey

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mdve

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mcl

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ey

gs

mb 50 µm

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lp

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lp

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ecm

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pec

mep

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pgc spg

pso

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pso

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cb

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mi

cb dis dis

bb 1 µm

pso

1 µm

7.4 Histological structure, cytology, and functional significance of organ systems 

diversification of processes (two at either side in P. nana and P.  gotoi), the gradual increase in their rigidity, the involvement of the tail fin in forming the processes, and, most importantly, the occurrence of two small appendicular muscles as well as four longitudinal strands of raising muscles. The appendicular muscles most likely arose from a slight dislocation and transformation of the former lateral secondary muscles, they are connected to the internal face of the epidermal ECM in immediate vicinity of the appendages (found in P. nana). In P. gotoi, the appendicular muscles are associated with the anterior process, so that the raising muscles in the tail are not the result of a transformation of muscle precursors and, thus, represent exclusive additions (an apomorphy) of this species (cf. Casanova et al. 2003).

7.4.1.8 Bioluminescent organs Judging from their position, cytoplasmic composition, and functional mechanism, the multicellular bioluminescent organs may be considered derivatives of the generally glandular distal epidermal cells. To date, bioluminescence has been observed in two meso- and bathypelagic chaetognath species only, namely Caecosagitta macrocephala and Eukrohnia fowleri (Haddock & Chase 1994; Thuesen et  al. 2010). In C. macrocephala, the bioluminescent organs are located at ventrolateral margin of either anterior lateral fin, whereas in E. fowleri a single bioluminescent organ is restricted to a circumferential band extending across the tail fin. In both species, each bioluminescent organ consists of clusters of huge granulated epithelial cells which typologically represent derived distal epidermal cells. Their cytoplasm is described to be compartmentalized into a honeycomb-like set of “membrane-bound chambers” each of which endowed with membrane-delimited, ovoid to elongated, and extremely electron-dense granules. The granules include densely packes “droplets” housed in a striated, paracrystalline

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matrix. According to the nomenclature introduced by Thuesen et al. (2010), it is assumed that these granules, termed as “bioluminescent particles”, function as compartments where the bioluminescence is produced. The chaetognaths’ ability to produce light is driven by a unique variant of the well-known luciferin-luciferase complex. Here, luciferase works in conjunction with coelenterazine as a to-be-oxidized substratum to emit light with maximum intensity at 467 nm wavelength (data obtained from C. macrocephala). As revealed by SEM observations, the bioluminescent compartments look hexagonal in shape, more or less obvious in C. macrocephala but the pattern is highly ordered in E. fowleri. Contents of the bioluminescent granules are released outside the fins’ surface in the event of mechanical stimulation. However, it is not clear yet whether the release of bioluminescent material is the result of apo- or merocrine secretion. Likewise, it is not exactly known where the oxidation of coelenterazine takes place, within or outside the granules. Thuesen et al. (2010) advocated for the latter option and assumed that upon being discharged onto the surface of the fins the membrane lining each granule may break open exposing its contents to either ions or oxygen triggering the or needed for the activation of the luciferin-binding protein. To date, bioluminescent organs seem to have evolved independently in the two phylogenetically distant chaetognath subclades Sagittidae and Eukrohniidae (Thuesen et al. 2010, Haddock et al. 2010, see also Gasmi et al. 2014 and fig. 7.8. in this book chapter). The independent acquisition of bioluminescence in deep sea-inhabiting chaetognaths may have derived from independent predation of luminous pelagic invertebrates, such as copepods, supplying the luciferin equivalent (Thuesen et al. 2010). The luciferase catalyzer, however, is assumed to be produced by the chaetognaths themselves (Haddock et al. 2010). The two so far recorded chaetognath species using bioluminescence may effectively deter and irritate predators or startle prey into swimming away and thus make it detectable to the ciliary fence and tuft organs (Thuesen et al. 2010).

◂ Fig. 7.21: Outer appearance (dissection microscopy: A) as well as histological (LM: B–C) and fine structural (TEM: D–G) anatomy of eyes in chaetognaths. A, Dorsal aspect of head of Spadella valsalinae in living state. B, Cross section of the posterior head region of Ferosagitta hispida, section level exemplarily indicated by dotted line in A. C, Overview of left eye of Spadella cephaloptera in cross section. D–G: Ultrastructural details of the eyes of S. cephaloptera. D, Cross section at lower magnification showing constitutive photoreceptor cells and central screening pigment cell (pgc), encapsulated by ECM of the epidermis (ecm). Somata of photoreceptor cells (pso) are located at the periphery, ciliary substructures of photoreceptor cells meet at the center enveloped by ramified screening pigment cell. E, Close-up of screening pigment cell and surrounding distal segments (dis) of photoreceptor cells; highly prismatic screening pigment granules (spg) are orderly aligned in parallel fashion. F, Close-up of distal segments containing complex apparatus of lamellae (la). G, Apices of two photoreceptor cells in longitudinal section showing the aberrant ciliary process consisting of the basal body (bb), the conical body (cb), and the endolamellar distal segment (shown in D–F). Further labels: at, anterior teeth; cc, corona ciliata; es, esophagus; ey, eye; gs, grasping spines; ho, hood; lp, lateral plate; mb, bilobate muscle (musculus bicornis); mcl, lateral muscle (musculus complexus lateralis); mdve, external vestibular dilator muscle (musculus dilator vestibuli externus); mep, multilayered epidermis; mi, mitochondrion; mos, mediolongitudinal-oblique head muscles (musculus obliquus superficialis); pec, proximal epidermal cells.

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7.4.2 Sense organs1 The epidermis of chaetognaths is rich in various kinds of ciliary sense organs showing different degrees of structural complexity. The highest diversity of sense organs is found on the head: the unpaired corona ciliata (Figs. 7.1 B, 7.2 B, 7.25, and 7.39 A) and the retrocerebral organ (Fig. 7.20) as well as the paired eyes (Figs. 7.2 A and 7.21) are present at the dorsal flank of the head and/or the transition zone between head and trunk. In addition, the head epidermis at least of S. cephaloptera was observed to include solitary or aggregated receptor cells (Fig.  7.19), being either scattered over the dorsal (presumably vibration-sensitive “receptors with large, non-motile cilia”: Horridge & Boulton 1967; Bone & Pulsford 1978; see also Fig.  7.19  B in this chapter) or ventral (presumably chemosensory “enclosed ciliary slit receptors”: Bone & Pulsford 1978) surface. In S. cephaloptera, these receptors protruding large cilia also occur on the tail. However, axons have not reported yet to project from these single receptor cells, so their sensory function is likely but not definite. Further small cephalic sense organs are located in three areas close by the mouth opening: (1) posterolateral to the mouth and hidden in the transvestibular pores (“posterolateral group”, see Figs. 7.5 A and 7.19 C–E), (2) (antero)lateral to the mouth associated with the vestibular ridges (“vestibular group”, see Fig.  7.5  A), and (3) immediately surrounding the mouth opening (“mouth group”) (Bone & Pulsford 1984; Thuesen et  al. 1988b). Spherical or spindle-shaped aggregates of receptor cells, called “ciliary fence organs” and “ciliary tuft organs” (Müller et al. 2014), are spread over the entire body in a specific pattern and classified as multifunctional sense organs (Figs.  7.24, 7.28  D, 7.31  C, and 7.39  A), probably most important to enable chaetognaths to be ambush predators in the sea (e.g., Feigenbaum & Reeve 1977; Feigenbaum 1978; Feigenbaum & Maris 1984; reviewed by Müller et  al. 2014). Besides the eyes, the definite functions of both uni- and multicellular sense organs are often not yet clear. Generally, all sense organs with stiff and exposed cilia have been affiliated with mechano(rheo)reception, whereas those with motile, internalized (corona ciliata), or strongly transformed (retrocerebral

1 The structural diversity of sense organs and their connection to the peripheral and central nervous system were previously and extensively reviewed by a team of authors (Perez et  al. 2014, Harzsch et al. 2016). The following section is in part reprinted from these recent contributions but also includes some additions of data and novel interpretations of particular sense organs and their putative function.

organ) cilia may perceive chemical cues (Shinn 1997; Müller et  al. 2014). Key papers dealing with the structure and putative functions of chaetognath sense organs to regard besides this review were provided by Hertwig (1880), Burfield (1927), Kuhl (1938), Ghirardelli (1968), Bone and Pulsford (1978, 1984), Goto and Yoshida (1984, 1987), Goto et  al. (1989), Bone and Goto (1991), Shinn (1997), Malakhov et  al. (2005), Müller et  al. (2014), and Harzsch et al. (2016).

7.4.2.1 Retrocerebral organ The retrocerebral organ is a bilobed, internalized epithelium of aberrant uniciliated receptor cells incorporated into the posterior domain of the brain (Figs. 7.20, 7.27 E, and 7.29 E). The receptor cells are assembled in two sacs (syn.: “cavities” in Malakhov & Frid 1984) in bilaterally symmetrical position relative to the mediosagittal plane of the brain (syn.: “globular masses” in Scharrer 1965) (Figs.  7.27  E and 7.29  E). Distally, both sacs taper into narrow canals (syn.: “retrocerebral canal” in Shinn 1997) that converge to a common canal close by the retrocerebral pore. Based on TEM examinations, previous authors noticed that the lumen of both sacs, the retrocerebral canals, as well as the inner space of the retrocerebral pore are filled with numerous, ramifying and intertwining cellular processes which they figured for microvilli (Scharrer 1985; Malakhov & Frid 1984). However, Shinn (1997) in part corrected this interpretation by stating that the “microvilli” contain microtubules and retrocerebral canal cells project highly modified cilia. By means of TEM data gained for this chapter, we can now confirm that each epithelial (receptor) cell lining and establishing the retrocerebral organ of S. cephaloptera projects a single but highly modified cilium (Fig. 7.20 C, E). A ciliary body and ciliary rootlet are observed at the apex of each receptor cell (Fig. 7.20 C, E). At the base of each ciliary process, the microtubules display a distinct axonemal 9 × 2 + 0 configuration; further distally, in the strongly ramified part, the microtubular pattern is unsorted (Fig.  7.20  D). Thin cytoplasmic processes are crammed into the narrow interspace of the glial-like sheath surrounding the entire retrocerebral organ and the somata of its receptor cells. These axon-like strands feed into a neurite bundle that penetrates the glial-like sheath at the ventrolateral face of the retrocerebral organ (Fig. 7.20 E) and finally enters the posterior (sensory) neuropil of the brain (Fig. 7.27 E). This may indicate the presence of primary receptor cells, although synaptic contacts or neuroendocrine vesicles could not be detected yet. Both sacs with modified ciliary

7.4 Histological structure, cytology, and functional significance of organ systems 

receptor cells taper distally into separate canals, which are lined by flattened epithelial cells projecting microvilli but lack cilia (Fig.  7.20  F). By invading the epidermis, both canals converge into a single one which passes through the epidermis and opens middorsally via the retrocerebral pore. Although a fine-scale analysis of the cellular organization in the retrocerebral organ of S. cephaloptera is still missing (Müller et al. unpublished data), the new TEM data presented here appear sufficient to strengthen previous ideas according to which the retrocerebral organ may be considered a proper sense organ. Scharrer (1965) saw structural correspondences to globular clusters of rhabdomeric receptor cells within the cerebral ganglion of the cladoceran crustacean Leptodora kindtii. Intracerebral ocelli of crustaceans are interpreted as accessory photoreceptors (e.g., Meyer-Rochow 2001; “brain photoreceptors” in Strauss & Dircksen 2010). However, the receptor organelles in the tognath retrocerebral organ are ciliary rather than chae­ rhabdomeric. Therefore, the scope of intra- or pericerebral sense organs to be compared and potentially homologized with the retrocerebral organ of Chaetognatha needs to be reassessed. Potential candidates for such a comparison may be the cerebral (“cephalic”) organs of Nemertea (compare Ling 1969, 1970; Beckers & von Döhren 2016) or multicellular (ad)cerebral organs of larval and adult polychaetous Annelida (nuchal organs: compare review of Purschke 1997; multicellular cerebral organs in Hausen 2007; Purschke 2005; Purschke et al. 2006; and Purschke 2016 for review) as these organs tightly adjoin the posterior aspect of the brain (“cerebral ganglion”) and exhibit an invaginated epithelium of ciliary receptor cells having access to the outer environment by a sensory canal and pore opening. Perhaps, the retrocerebral organ and its cellular constituents may provide a promising substrate for allying chaetognaths to lophotrochozoans. The above-mentioned organs of nemerteans and annelids are proposed to serve as chemoreceptors, neuroendocrine organs, or accessory photoreceptors (e.g., Ling 1969, 1970; Ferraris 1985; Purschke 1997, 2016). Therefore, these functions should be also taken into consideration when discussing the putative function(s) of the retrocerebral organ of chaetognaths. Shinn (1997) previously proposed that it might be a baroreceptor.

7.4.2.2 Eyes The structural diversity of chaetognath eyes has not been fully explored yet, but the available studies indicate significant taxon-specific variations of eye design (Ducret 1975, 1977, 1978; Goto & Yoshida 1981, 1983, 1984;

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Goto et al. 1989; reviews Bone & Goto 1991; Shinn 1997; Perez et  al. 2014; Harzsch et  al. 2016). The photoreceptors of chaeto­gnath eyes bear a highly modified cilium (Fig.  7.21  D–G). They are dorsally located in the epidermis at either side of the head (Fig. 7.21 A), where they are encapsulated by the epidermal ECM and several sheath cells (Fig. 7.21 C). Based on the orientation of the photoreceptor cells, chaetognath eyes are classified as “indirect” eyes in the Sagittidae (including P. draco) and Spadellidae (Fig. 7.22), and as “direct” eyes in the Eukrohniidae (Fig. 7.23). The well-studied “indirect” (syn.: “inverted”) eyes exhibit one central pigment cell that is surrounded by 70 to 600 photoreceptors cells per eye depending on the species (Figs. 7.21 and 7.22). The photoreceptor cell somata are located in the unpigmented peripheral region of the eye. Their axons project into the optic nerve toward the posterior compartment of the brain (Rieger et al. 2010; see also Figs. 7.21 D and 7.22 C in this chapter). They extend a receptoral process toward the single pigment cell, the so-called distal segment, which most likely contains the rhodopsin-like photopigments (Goto & Yoshida 1988). The distal segment is anchored in an axial centriole by a connecting piece with a 9 × 2 + 0 microtubular configuration so that the entire receptoral process was identified as a highly modified cilium (Shinn 1997) (Figs. 7.21 G and 7.22 B). The “conical body” of the receptoral process (Figs. 7.21 G and 7.22 B–C) adjoining the distal segment proximally is considered unique among metazoan photoreceptor cells (Eakin & Westfall 1964; Goto et al. 1984). It has a clear and refractive appearance in live animals so that a dioptric function was discussed. Ultrastructural studies showed that its proximal part is composed of dense granules (Figs.  7.21  G and 7.22  B), whereas its distal part contains irregular cords of undefined material (Eakin & Westfall 1964). Goto et al. (1984) discussed a possible role as participating in the organization of the photoreceptor membranes in the distal segment. These are arranged as lamellar stacks, which enclose a 30- to 45-nm-wide intralamellar lumen (Figs.  7.21  E–F and 7.22  A). Mediany, distal segments adjoin the screening pigment cell showing a highly ordered alignment of polymorphic but extremely electron-dense pigment granules. In S. cephaloptera, these screening pigment granules strictly display a rectangular shape (Fig. 7.21 D, E). Within distal bodies apposing membranes of a lamella are continuous across this space at numerous pores. These pores have an 80- to 95-nm center-tocenter distance and are regularly aligned in a grid-like pattern of almost crystalline consistency. The lamellae are separated by 10–20 nm of interlamellar cytoplasm

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Fig. 7.22: Indirect visual system of chaetognath. A, Schematic representation of the distal lamellar structure of a visual photoreceptor cell from the eye of Paraspadella gotoi (previously referred as Spadella schizoptera, after Goto et al. 1984). B, Photoreceptor cell from the eye of Pseudosagitta scrippsae (after Eakin & Westfall 1964). C, Diagram of a cross section through an indirect chaetognath eye (after Shinn 1997). Modified and reprinted from Perez et al. (2014).

(Fig. 7.22 A). The cytological origin of the stacks of perforated lamellae is unclear with respect to the conical body. Goto et  al. (1984) concluded that such lamellar arrays are not present in any other bilaterian photosystem so that we interpret the entire ciliary photoreceptive apparatus as another autapomorphy of this taxon. Goto et al. (1989) comparatively analyzed the eye structure in 10 sagittid species. The size and shape of the pigment granules were highly variable, as was the arrangement

of the photoreceptive region. On the basis of these variations, they defined five different types of indirect eyes and showed that this classification may be related to the depth of habitat. In Eukrohnia, the apices of the photoreceptor cells are directed distally toward the body surface and incoming light (Fig. 7.23 D). These “direct” eyes appear to have dozens of closely packed hexagonal facets (conical bodies = “corps coniques”), which represent crystalline-like,

7.4 Histological structure, cytology, and functional significance of organ systems 

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Fig. 7.23: Direct visual system of chaetognath. A (light micrograph), B (SEM micrograph)—Dorsal view of the eye of Eukrohnia hamata showing the anterior (arrow) and posterior (arrowhead) group of facet-like ommatidia (after Ducret 1975). C, Photoreceptor cell from the eye of E. hamata (drawn from ultrastructural observations of Ducret 1975, 1978). D, Diagram of a cross section through a direct chaetognath eye. The retinal bodies are sectioned longitudinally (drawn from ultrastructural observations of Ducret 1975, 1978; Bone & Goto 1991).

condensed apical parts of the likewise strongly modified photoreceptive cilia (Fig.  7.23  A–B). Ducret (1975, 1977, 1978) termed these facets “ommatidia” because they resemble corneal facets of arthropod ommatidia (Fig.  7.23  A–B). In Eukrohnia species belonging to the hamata group, the photoreceptor cells develop a complex array of apical microvilli that surrounds the centralized

conical body. Because of their highly ordered appearance, Ducret (1975, 1977, 1978) considered these microvilli to be comparable with rhabdomeres (“rhabdomères”), similar to those found in arthropod ommatidia, and thus seemed to have identified a further, potentially light-sensitive part of the cell (Fig. 7.23 C). To date, it remains uncertain whether the modified cilium and/or the surrounding

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microvilli are true photoreceptor organelles. The number of facet-like ommatidia decreases as a function of depth, similar to the disappearance of photopigments in species living in obscure habitats such as deep-sea or cave environments, for instance, in Heterokrohniidae (Casanova 1986a, 1986b, 1992; Thuesen & Haddock 2013), Eukrohniidae (Casanova 1986c), and Spadellidae (Bowman & Bieri 1989; Casanova 1996). There is considerable variation in the architecture of the photoreceptive membranes, and in other Eukrohnia species (fowleri group), the eye is not facetted (Ducret 1975, 1977; Bone & Goto 1991). As it has been the case during the diversification of the muscular system (Casanova & Duvert 2000; see Section 7.4.4 in this chapter, “Muscular Apparatus and Locomotion”), the Chaeto­gnatha seem to have thoroughly “experimented” with eye design at the histological and cytological levels during their long and isolated evolutionary pathway.

7.4.2.3 Ciliary fence and tuft organs Additional sense organs used for orientation, prey detection, and predator avoidance are located in the epidermis of the chaetognath head. Other, structurally less complex ciliary sense organs are the peristomatic, presumably chemosensory “enclosed ciliary slit receptors” (Bone & Pulsford 1978) on the head as well as “large cilia” extended by single sensory cells, which occur in various positions of the body and are assumed to function as vibration receptors (Horridge & Boulton 1967; Bone & Pulsford 1978) (Fig. 7.19 B). Chaetognaths are also equipped with three more types of ciliary sense organs (Malakhov et al. 2005; Müller et al. 2014): the transversally oriented ciliary fence organs (Figs. 7.24 A–D, F–G, 7.31 C, 7.33 D, and 7.39 A), the longitudinally (parallel

to the anterior-posterior axis) oriented ciliary tuft organs (Figs. 7.24 A, E, H and 7.28 D), and a ciliary loop, the corona ciliata (Figs. 7.1 B, 7.2 B, 7.24 A, 7.25, and 7.39 A). The ciliary fence and ciliary tuft organs (sensu Malakhov et  al. 2005) are conspicuous multicellular sense organs spread over the head (e.g., Figs. 7.28 D and 7.31 C), trunk (Fig.  7.39  A), and tail, well visible by a fan of stiff bristles also on a living specimen observed through a dissection or light microscope. Hence, these organs were termed as “bristles” or “hairs fans”, but are also known under the synonyms “sensory cells”, “ciliary sensory organs”, “ciliary fence organs”, “ciliary fence receptors”, or just “fence receptors” (Bone & Pulsford 1978; Feigenbaum 1978; Nagasawa & Marumo 1982; Welsch & Storch 1983a; Shinn 1997). Shape, number, and distribution of the ciliary fence organs are species-specific and can be easily highlighted by immunohistochemical techniques making these a useful tool for taxonomic studies (Müller et  al. 2014). Several authors have studied the outer and internal ultrastructure of ciliary fence and ciliary tuft organs utilizing electron microscopic techniques, such as SEM and TEM (Horridge & Boulton 1967; Bone & Pulsford 1978; Welsch & Storch 1983a; Ahnelt 1984; Bone & Pulsford 1984; Goto & Yoshida 1987; Shinn 1997; Malakhov et al. 2005; Müller et al. 2014). Inspection of SEM images is, however, deceptive because the receptoral cilia have lost their rigidity and look twisted (Fig.  7.24  A–B). The “wooly” appearance of cilia is probably a fixation and/or dehydration artifact. Ciliary fence and tuft organs are common and embedded in the epidermis where they form globular or diamond-shaped aggregates of receptor and sheath cells (e.g., Fig. 7.39 A). Their arrangement in the epidermis differs relative to the body and with respect to several ultrastructural characters (Müller et al. 2014): ciliary lines of ciliary fence

▸ Fig. 7.24: Outer and inner morphology of ciliary fence and tuft organs in various chaetognaths: Parasagitta setosa (A–B), Spadella cephaloptera (C–D, F–H), Ferosagitta hispida (E). SEM: A–B; TEM: C–H. Originals. A, Neck region showing anterior half of corona ciliata (cc) in mediolongitudinal position. Ciliary fence organs (cfo) have their protruded cilia oriented perpendicular to body’s length axis, whereas in ciliary tuft organs (cto) cilia are aligned parallel to it. B, Close-up of ciliary fence organ as seen in A (dotted rectangle), rigidity of receptoral cilia (ci) is lost due to chemical fixation and/or SEM preparations. C, Cross-cut ciliary fence organ located on the dorsal midline of the trunk, central type 1 receptor cells (rc1) are bidirectionally framed by multiple lines of type 2 receptor cells (rc2). D, High-power magnification of ciliary insertion area of a ciliary fence organ; central depression where apices of both types of receptor cell meet, each receptor cell protrudes a sensory cilium, type 1 cilia (ci1) show a bigger diameter than type 2 ones (ci2), magnified sector is indicated by dotted rectangle in C. E, Close-up of apical portions of type 2 receptor cells of a paramedian ciliary tuft organ on dorsal trunk hemisphere; note the elongated, cross-striated ciliary root portion (csr) projecting from extremely electron-dense distal root body (edr). F, Longitudinal section of apical region of a dorsolateral ciliary fence organ. Several type 1 receptor cells are cut; the distal, electron-dense portion of the ciliary rootlet is elongated and lacks a subjacent, cross-striated part. G, Close-up of apices of type 1 receptor cells in longitudinal section close to level shown in F; note lack of cross-striated portion of ciliary rootlet and typical sequence of adhering junctions. H, Horizontal section of the apices of type 1 and type 2 receptor cells of a dorsomedially positioned ciliary tuft organ. Note the decussate formation of filamentous bundles (fis) attaching the distal, electron-dense rootlet portion to the apical cell membrane. Further labels: aj, desmosome-like adhering junction; bb, basal (ciliary) body; dec, distal epidermal cell(s); ecm, extracellular matrix; h, head; me, mesenterium; mi, mitochondrion; mic, microtubules; myc, myoepithelial cells; pec, proximal epidermal cells; sej, septate junction; sg, secretory granule; shc, glandular sheath cell; ssg, secretory granule of sheath cell; stmu, paramedian secondary musculature; tr, trunk; v, vacuole.

7.4 Histological structure, cytology, and functional significance of organ systems 

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Fig. 7.25: Ultrastructure of the corona ciliata of Spadella cephaloptera as revealed by SEM (A) and TEM (B–I). Originals. A, Overview of corona ciliata, dorsal view. B–E: Longitudinal sections demonstrating the ultrastructural architecture of the corona ciliata. B, Multiple image alignment of the right half of the corona ciliata giving an overview of the distribution of the four cell types involved in its construction. C, High-power magnification of the apical region of putative absorptive cells and receptor cells displaying an enormous microtubular apparatus. The apices surround a subsurficial cavity narrowed by partly ramified microvilli-form projections and cilia (not shown!). D, Close-up of the apical region of the outer cellular portion, which forms a highly ordered system of multilayered, narrowed, collar-like tips from which the vibratile cilia

7.4 Histological structure, cytology, and functional significance of organ systems 

organs are aligned transversally, perpendicular to the mediosagittal plane, whereas they are oriented parallel to the mediosagittal plane in ciliary tuft organs. Ciliary fence and tuft organs contain numerous monociliary receptor cells (Fig. 7.24 C–D, E–G) and were discussed as either representing a primary (Bone & Pulsford 1978) or a secondary type of receptor (Reisinger 1969; Welsch & Storch 1983a). Shinn (1997) suggested that reports according to which the fence receptors are secondary sensory cells were more reliable. In S. cephaloptera and S. valsalinae, two different types of multilayered and partly pooled primary receptor cells constitute the ciliary fence and ciliary tuft organs, named type 1 (Fig. 7.24 C, F, H) and type 2 receptor cells (Fig. 7.24 C–D, H) (reinvestigated by Müller et al. 2014). Both types of receptor cells extend a single, nonlocomotory cilium from the narrow apex so that multiple rows of highly ordered cilia are formed (Figs. 7.24 C–H and 7.33 D). However, their location is different. Type 1 receptor cells form a single line along the midline axis of the organ, whereas type 2 receptor cells surround the type 1 cells in multiple rows (Fig. 7.24 C, H). There are also ultrastructural differences. Each type 1 receptor cell extends a long and thick cilium (Fig. 7.24 D, F), its ciliary rootlet is undivided but sac-like and jagged (Fig.  7.24  D, F–H). By contrast, the thinner and slightly shorter cilium of a type 2 receptor cell is strengthened and/or adjusted by a bipartite ciliary rootlet consisting of an almost amorphous and highly electron-dense distal portion and a cross-striated proximal portion, which often reaches deeply into the receptor cell soma (Fig. 7.24 D–E, H). These type-specific ultrastructures of the ciliary apparatus may vary also among ciliary fence and ciliary tuft organs. It is known that the ciliary fence and ciliary tuft organs of chaetognaths which are abundant in large numbers across their entire body innervate the ventral nerve center (e.g., Hertwig 1880;

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Bone & Pulsford 1984). Experiments showed that the receptor cells in the ciliary fence and ciliary tuft organs detect hydrodynamic stimuli and react to close-range mechanosensory input and are used to initiate attack or escape movements (Horridge & Boulton 1967; Feigenbaum & Reeve 1977; Feigenbaum & Maris 1984; Bone & Goto 1991). Specific components of the basal apparatus (proximal to the basal body) in type 1 and type 2 receptor cells of ciliary fence and tuft organs are unique among ciliary receptors of metazoans, namely, the extremely electrondense body that is connected to belt-like adhering junctions by numerous microfilaments (Müller et  al. 2014; see also Fig.  7.24  E, G–H in this chapter). Consequently, this feature, along with ciliary fence and tuft organs as a whole, has to be added to the extensive list of chaetognath apomorphies. The peculiar configuration of the basal apparatus may be involved in the differential transmission of various mechanical stimuli or, alternatively, ensure reshaping of the basal body in the cylindrical apex of the receptor cell in case the receptive cilium is bent and a receptor potential is released (Müller et al. 2014).

7.4.2.4 Corona ciliata The corona ciliata is located on the dorsal side of the body and extends along the dorsoposterior neck region (Figs. 7.1 B, 7.2 B, 7.21 B, 7.24 A, 7.25 A, 7.29 A, and 7.39 A) (Ghirardelli 1959; Bone & Goto 1991; Shinn 1997). It consists of numerous monociliary cells forming a loop and shows an extensive and intricate pattern of innervation. The ultrastructure of the corona ciliata has been described in different chaetognath species by several authors (Horridge & Boulton 1967; Nagasawa & Marumo

◂ are protruded. The microtubular system of the ciliary epithelial cells becomes considerably condensed. Note the profiles of meshwork of extracellular canals (mwc) connecting lumen of subsurficial cavity to the invaginated collar around bases of vibratile cilia (arrows) filled with environmental water. E, Details of the apical region of a ciliary epithelial cell cut at the level of the basal body and distal portion of the ciliary rootlet encompassed by microtubules; arrows point at invaginated, periaxonemal space filled with environmental water. F–I, Innervation of the corona ciliata following a gradient of structural complexity starting from (F) primary axon bundles formed by several local sensory cells of the inner cellular portion, which (G) become collected in lateral branches (H) passing into and subsequently forming both coronal nerves. I, The coronal nerve may contain both afferent and efferent fibers. At the ventral margin of each coronal nerve, several synapses are found that may innervate the paired dorsomedian longitudinal musculature. Further labels: abc, absorptive cells; axt, axonal terminal; bci, basal region of the locomotory cilia (surrounded by cytoplasmic collars); bcn, lateral branches of a coronal nerve; c, locomotory cilia; ca, subsurficial cavity (filled with cilia and microvilli); cb, ciliary (basal) body; cc, corona ciliata; cn, coronal nerve; co, collar-like infolding of the ciliary epithelial cells; cplc, epithelial cells producing locomotory cilia; cr, ciliary rootlet; ctu, cytoplasmic tubules; dep, distal epidermal cell; dlmu, paired dorsomedian longitudinal musculature; ecm, extracellular matrix (of the epidermis); end (primary) endosome; h, head; ip, inner cellular portion of the corona ciliata; ltmu, primary longitudinal (trunk) musculature; mcib, multiple and highly ordered bands of locomotory cilia; mi, mitochondrion (tubular type); mic, microtubules; miv, microvilli-form apical projections of the putative absorptive cells and sensory cells; n, nucleus (absorptive cell); ne, neck-like lateral protuberance at the head-trunk transition zone (collarette region); neu, neurites; nt, neurotubules; op, outer cellular portion of the corona ciliata; pab, primary axon bundle of coronal sensory cells; pep, proximal epidermal cells; sc, sensory cell; shc, sheath cell; syv, synaptic vesicles; tr, trunk.

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1982; Malakhov & Frid 1984; Shinn 1997; Giulianini et al. 1999; Malakhov et al. 2005). The macroscopic shape and orientation of this organ were suggested to be species specific (Tokioka 1965a, 1965b; Bieri 1991b; Shinn 1997), although some intraspecific variations of its size, shape, and position on the body may occur in relation to age and zoogeographic distribution (Ghirardelli 1968). Many of the constitutive cells in the corona ciliata of adult specimens of S. cephaloptera show mitotic activity suggesting a continuous turnover of cells (Müller et al. 2014). Shinn (1997) described the ultrastructure of the corona ciliata of the F. hispida. In his semischematic reconstruction (Fig. 7.53, p. 163), he revealed the existence of an outer portion of corona-constituting cells (“lateral cells”), represented by a multilayer of receptor cells forming short and vibratile cilia, and an inner portion of cells (“medial cells”), characterized by functional or degenerating secretory cells which also form a single cilium. The cilia of the secretory cells are not visible from outside as they extend into a subsurface canal that is in contact with a meshwork of extracellular spaces around the apices of the cells of the outer portion, and subsequently, to the outer environment via the collar-cilium interface. The secretory cells of the corona ciliata were assumed to release an electron-dense secretion into the subsurface cavity, whereas Ghirardelli (1968) reported that corona-associated gland cells produce secretion, which spreads midposteriorly along the dorsal midline and flows further posteriorly by turning to lateral flanks of the trunk where it gets finally accumulated at and within the female gonopores. Because of the continuous activity of ciliary motion, ongoing secretion, and persistent cell divisions, Shinn (1997) suggested that coronal epithelial cells are short-lived and proceed through sequential changes in morphology. The general subdivision of corona ciliata epithelial cells in an outer and inner portion, as depicted by Shinn (1997) in F. hispida, is also found in S. cephaloptera (Fig. 7.25 B). New TEM observations, in part first published in this chapter, indicate the existence of two distinct cell types in the inner portion of coronal epithelial cells, namely, absorptive cells projecting two or several cilia and receptor cells having a single cilium (Fig.  7.25  C). Secretory cells are restricted to the periphery of the corona ciliata and represent elongated distal epidermal cells (Fig.  7.25  B, D). Cilia of absorptive cells and receptor cells are protruded in the small subsurface cavity, which is also riddled with microvillar processes and electron-lucent spherules (Fig. 7.25 B–C). A meshwork of tiny extracellular canals connects the deeply invaginated extracellular collar around vibratile cilia of outer epithelial cells, carrying environmental water, with the lumen of the subsurficial “receptor cavity” (Fig. 7.25 D–E). The putative

absorptive cells resemble midgut epithelial cells by carrying cytoplasmic tubules, numerous vacuoles with heterogenous content (probably endosomes), and a huge centroproximal lysosome. These cells likely modify the quality of the subsurficial cavity water by absorbing unfit chemicals. Thereby, sensitivity of the receptor cells may be enhanced. The axons of the primary receptor cells become bundled at the medioproximal periphery of the inner cellular portion (Fig.  7.25  F) and project toward the center of the corona ciliata (Fig. 7.25 G) where they converge and pass into the dorsomedian coronal nerves that house approximately 400 neurites and project into the posterior area of the brain (Müller et  al. 2014; see also Fig.  7.25  H in this chapter). Whether or not the corona really contains primary or secondary receptor cells has long been a matter of dispute (Bone & Pulsford 1984; Malakhov & Frid 1984), but in his drawing, Shinn (1997: fig. 53, p. 163) hints at the presence of neurites basal to the outer margin of the coronal cell cluster. New immunohistochemical and TEM data, however, indicate that synapses are found throughout the coronal nerves, close to the ECM and the subjacent longitudinal dorsomedian musculature of the trunk (Fig. 7.25 I). The function of the corona ciliata is not yet known, but the new insights into the microscopic architecture (Müller et al. in preparation) support previous assumptions in favor of chemoreception. Volumetric measurements (visualized using PIV= particle image velocimetry) of water flow created by the beating cilia of the corona in S. cephaloptera (Bleich et al. 2017) additionally support the assumption of a sensory function. Indeed, seawater is accelerated toward the corona ciliata from anterior of the body in a funnel-shaped pattern and expelled laterally and caudally, with part of the water being recirculated. Although the corona ciliata is well suited to work as a multifunctional organ, including secretion, respiration, and excretion, chemoreception appears likely to be the main function. The funnel-shaped directional flow can possibly enable directional chemosensation, and olfactory sensitivity may be enhanced by resampling the same volume of seawater (Bleich et al. 2017). The evolutionary origin of the corona ciliata still needs to be evaluated. Previous attempts have failed to find an equivalent multicellular ciliary sense organ with direct innervation from the brain in other bilaterian taxa. The retrocerebral organ (see previous section) and the nuchal organ of annelids, and among them especially those equipped with an “olfactory chamber” (compare Fig. 7.13 in Purschke 1997), may be interesting objects of fine-scale multimodal microscopic comparison. It would not be utterly surprising if the corona ciliata turns out to be among the oldest olfactory organs in the stem lineage of the Bilateria.

7.4 Histological structure, cytology, and functional significance of organ systems 

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Fig. 7.26: General organization of the chaetognath nervous system. A, Schematic diagram (modified from Kapp 2002); B, confocal laserscan microscopy. Sagitta bipunctata, obliquely ventral view of the ventral nerve center, immunohistochemical labeling of tyrosinated tubulin (red), and histochemical labeling of nuclei (blue). Reprinted from Perez et al. (2014).

7.4.3 Nervous system² The nervous system of chaetognaths comprises peripheral and central components of equal complexity (Fig. 7.26). The central nervous system is mainly situated in the head housing the (1) dorsally located brain (syn.: “cerebral ganglion” sensu Goto & Yoshida 1987; see Figs. 7.27 and 7.29) including the retrocerebral organ (see previous section and Fig.  7.20) and the (2) deeply sunk circumesophageal chain of cephalic ganglia integrating the paired vestibular and (occasionally) esophageal (= “pharyngeal”) ganglia as well as the unpaired subesophageal ganglion (Figs.  7.26–7.28). Connectives and, most posteriorly, a subesophageal commissure

2 The architecture of the nervous system and sense organs was extensively reviewed by the team of authors (Perez et al. 2014; Harzsch et al. 2016). This section is in part reprinted from these recent contributions.

link these ganglia. The esophageal and subesophageal ganglia do not fit a strict definition of the term ganglion because they lack a core of synaptic neuropil and exclusively consist of neuronal somata (Goto & Yoshida 1987). The brain also connects to the ventral nerve center in the trunk via paired main connectives. The peripheral nervous system is complex and contains in part orthogonally arranged networks of single neurites or neurite bundles, called the intra- and basiepidermal neuronal plexus that makes contact to mechanoreceptors distributed over the entire body (see Fig. 7.33 and next section for details). The nervous system of Chaetognatha was studied with a variety of morphoanalytical methods comprising observations on dissected or whole-mounted specimens (e.g., Krohn 1844; Grassi 1883) and analyses utilizing classic histology (e.g., Hertwig 1880; Grassi 1883; Ritter-Záhony 1911; Burfield 1927; John 1933; Kuhl 1938; Bone & Pulsford 1984; Goto & Yoshida 1987), immunohistochemistry (Bone

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7.4 Histological structure, cytology, and functional significance of organ systems 

et al. 1987b; Goto et al. 1992; Duvert et al. 1997; Harzsch & Müller 2007; Harzsch et al. 2009; Rieger et al. 2010), and transmission electron microscopy (Ahnelt 1980, 1984; Bone & Pulsford 1984; Rehkämper & Welsch 1985; SalviniPlawen 1988; Harzsch et al. 2009). Most constituents of the central nervous system are strictly basiepidermal as they constantly rest on the epidermal ECM. The brain builds no exception from that rule as it fully rests on the ECM but is also tightly enwrapped by a multilayered system of extremely flattened, most basally located epidermal cells (e.g., Fig. 7.29 B, E–G). By contrast, the deeply sunk components of the circumesophageal chain of cephalic ganglia are entirely encapsulated by invaginated strands of the epidermal ECM (Fig. 7.28 B, E–F), however very thin in some places (Goto & Yoshida 1987; Shinn 1997). Only around the vestibular and esophageal ganglia the epidermal ECM sheath may vanish locally (Fig. 7.28 B–C). The nervous system is the first fully differentiated and operative organ system during chaetognath development (Rieger et  al. 2011; see Section 7.5.4, “Neurogenesis”). Besides this chapter, comprehensive summaries of the organization of the nervous system in chaetognaths, worthwhile to read not least in a historical context, were provided by Hertwig (1880), Burfield (1927), Kuhl (1938), Bullock (1965), Bone and Goto (1991), Shinn (1997), and Harzsch et al. (2016).

7.4.3.1 Brain and circumesophageal chain of cephalic ganglia The brain consists of a core neuropil with numerous synapses (Fig.  7.29  D) surrounded by clusters of neuronal somata (see Figs.  7.27  E and 7.29  B–C, E in this chapter; compare Goto & Yoshida 1987; Bone & Goto 1991; Rieger et al. 2010), and its fine structure has received much attention (Scharrer 1965; Ahnelt 1980, 1984; Rehkämper & Welsch 1985; Goto & Yoshida 1987; Salvini-Plawen 1988). It is situated immediately at the base of the cephalic pluristratified (multilayered) epidermis in a basiepidermal position resting on the epidermal ECM and encapsulated by specialized sheath cells representing extremely flattened proximal epidermal cells (Rehkämper & Welsch 1985; Salvini-Plawen 1988; Bone & Goto 1991; Shinn 1997)

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(Figs. 7.19 A, 7.20 A, and 7.29 A–B, E). At the anterolateral corners of the brain, two frontal connectives project ventrally toward the mouth region connecting with the circumesophageal chain of cephalic ganglia (Figs. 7.26 A and 7.27 A–D). Many bilaterally symmetrically arranged pairs of nerves project from particular cephalic ganglia toward different effector organs in the head and trunk (general overview provided in 3D models on Fig. 7.27 A–D), among them the frontal, dorsal (see also Fig. 7.28 D–E), vestibular, and mandibular nerves (projecting from vestibular ganglia) as well as the unpaired stomatogastric nerve that projects posteriorly from the subesophageal ganglion and is attached along the ventral midline of the fore- and midgut (Fig.  7.31  A–B). In S. cephaloptera, the stomatogastric ramifies into small, partly myelinated axon profiles that encompass the intestine. The myelinated axons directly adhere to the visceral ECM (VECM) in places where the line of specialized peri-intestinal muscle cells is interrupted. Peritoneocytes may enwrap the myelinated fibers and separate them from the trunk coelom (Fig. 7.40 F). The esophageal ganglion gives rise to the medially directed ventral esophageal nerve that seems to target the esophageal muscles (Shinn 1997). The vestibular ganglia are the largest among circumesophageal ganglia. The cross shape of this ganglion changes from ovoid (Fig. 7.28 A) to cone-like (Fig.  7.28  B). The neuronal somata flank the central neuropil dorsally, laterally, and ventrally (Fig.  7.28  B). The ganglion is incompletely wrapped by the epidermal ECM, and gaps may be found along the median rim (Fig.  7.28  B–C). The vestibular ganglia presumably control the operation of the grasping spines and receive sensory input from the vestibular ridge papillae surrounding the mouth via the frontal nerves (e.g., John 1933). Except for the elaborate intra- and basiepidermal neuronal plexus (see next section), all components of the cephalic nervous system are deeply sunk into the musculature that moves the grasping spines and the esophagus. Clusters of giant fibers are observed in the frontal connectives, circumesophageal commissure (Fig. 7.28 F), various peripheral nerves projecting from vestibular and esophageal ganglia (e.g., dorsal nerve: Fig.  7.28  E), and stomatogastric nerve (Fig. 7.31 A). Giant fibers are multiply enveloped by processes of glial cells (Fig. 7.28 F).

◂ Fig. 7.27: Cephalic nervous system of chaetognaths. A–D, 3D reconstruction in Ferosagitta hispida based on a series of approximately 400 transverse semithin sections through the head; structures of the nervous system are colored yellow, the gut is stained green. A, frontodorsal view; B, lateral view; C, frontoventral view; D, posterolateral view; Abbreviations: B, brain; CN, coronal nerve; DN, dorsal nerve; EY, eye; FC, frontal connective; FN, frontal nerve; MC, main connective; MN, mandibular nerve; OE, esophagus; OEC, esophageal commissure; ON, optic nerve; STN, stomatogastric nerve; VG, vestibular ganglion; E, confocal laser-scan microscopy. Components of the cephalic nervous system of Spadella cephaloptera. Immunolocalization of tyrosinated tubulin (red) and histochemical labeling of nuclei (blue). Reprinted from Perez et al. (2014).

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7.4 Histological structure, cytology, and functional significance of organ systems 

A pair of main connectives is projected to the side of the brain (Figs. 7.26, 7.27, and 7.31 C–D). The main connectives establish a connection between the brain and the ventral nerve center in the trunk (see Fig. 7.26 and description below). Thus, a second (posterior) ring around the foregut is formed. Peripheral organs associated with the brain are a pair of eyes (Goto & Yoshida 1984, 1988; Goto et al. 1984; 1989; see also Figs. 7.21, 7.26 A, and 7.27 in this chapter); the corona ciliata, which is localized in the dorsal part of the neck (Bone & Goto 1991; Shinn 1997; Müller et  al. 2014; see also Figs.  7.25, 7.26  A, and 7.27); and, nested within the posterior domain of the brain, the retrocerebral organ, a bilobed structure with an unknown putative sensory function, possibly a baroor chemoreceptor (see, for instance, Fig. 7.29 B, E; Section 7.4.2.1, “Retrocerebral Organ”; and Shinn 1997). Mediany, between both receptor cavities of the retrocerebral organ, profiles of each two optic and coronal nerves are located projecting posteriorly, immediately adjacent to the epidermal ECM (Fig. 7.29 E–G). The brain consists of a central neuropil divided into an anterior and a posterior domain flanked laterally by aggregated neuronal somata (Figs. 7.27 E and 7.29 B–C, E). Brain neuropil is extraordinarily rich in synaptic contacts and neuroendocrine terminals (Fig. 7.29 D). Few neuronal somata are also encountered at the interface of the central neuropil and the overlaying epidermis (Goto & Yoshida 1987; Shinn 1997). Fine structural analyses of the brain of P. setosa and F. hexaptera revealed that the sheath around the brain is composed of two distinct zones: the (1) outer zone shows flattened somata of the basalmost (proximal) epidermal cells that taper laterally into small projections, whereas the (2) multilayered inner zone lacks somata but instead shows numerous, extremely flattened, filamentous, and electron-dense cellular projections almost free

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of cytoplasmic organelles (Rehkämper & Welsch 1985; Goto & Yoshida 1987; Bone & Goto 1991). Rehkämper and Welsch (1985) emphasized close resemblance to myelin sheaths wrapping axons of vertebrate neurons. Like in the ventral nerve center (see Harzsch et al. 2009; and Section 7.4.2.6 in this chapter), the brain rests on the epidermal ECM. In addition, the brain is lined dorsally by tight sheath of multiple processes of most proximal epidermal cells, termed here as sheath cells. Besides their flat structure, sheath cells can be also distinguished from overlaying regular proximal epidermal cells by their extremely electron-dense contents (Figs. 7.20 B and 7.29 B, E). These flattened sheath cells also separates both lateral assemblies of neuronal somata into subordinate clusters by extending radial processes into the brain (Rehkämper & Welsch 1985; Goto & Yoshida 1987). These radial septa also isolate the central neuropil from the somata. However, there has not been an attempt yet to unravel the diversity of neuronal cell types, as based on TEM analyses, as it was done recently for the ventral nerve center (for details, see Section 7.4.3.2 in this chapter; and Harzsch et  al. 2009). Close similarities in topology and compartmentalization of neuronal somata as caused by the glial-like sheath cells make it reasonable to assume that the brain is organized according to constructional and functional principles comparable with those documented in the ventral nerve center (Harzsch et al. 2009). The distribution of neuroactive substances in the brain such as the biogenic amine serotonin and RFamidelike neuropeptides has been examined (Bone et al. 1987b; Goto & Yoshida 1987; Goto et al. 1992; Rieger et al. 2010) as well as aspartate immunoreactivity (Duvert et  al. 1997). Nerves emerging from the brain target esophageal and head muscles are described further below (see Section 7.4.4. “Muscular Apparatus and Locomotion”).

◂ Fig. 7.28: Ultrastructural aspects of subepidermal, circumesophageal ganglia and efferent nerves. A, D, Original light (LM) micrographs; B–C, E–F, Orginal TEM micrographs. A, Cross section of medioposterior head region of Ferosagitta hispida. B, Cross section providing overview of the arrangement of neuronal somata (nso) and central vestibular neuropil (vnp) of right vestibular ganglion (vsg) of Spadella cephaloptera; section level exemplarily indicated in A (dotted rectangle); note the incomplete coverage by the epidermal extracellular matrix (ecm), which is well visible at dorsal, lateral, and ventral borders of vestibular ganglion (arrows) but missed locally at its median rim (arrowheads). C, Close-up of centromedian rim of same vestibular ganglion as shown in B (dotted rectangle); boundary to surrounding head muscle lacks an extracellular matrix (indicated by arrowheads); synaptic contacts are frequently observed in vestibular neuropil. S. cephaloptera. D, Cross section of medioposterior head region of F. hispida, slightly posterior to A. E, Cross section of dorsal nerve containing numerous regular neurites (neu) and some profiles of giant neurons (gn) wrapped by projections of multiple glial sheath cells (gls); section level exemplarily shown in D (dotted rectangle). S. cephaloptera. F, Cross section of circumesophageal commissure (csec) of S. cephaloptera; indicated exemplarily by dotted rectangle in D; note the cluster of giant neuron fibers making connection with the stomatogastric system. Further labels: cc, corona ciliata; cemu, circumesophageal muscles; con, coronal nerve(s); cto, ciliary tuft organ; cu, cuticle; den, dendrites; es, esophagus, gs, grasping spines; hco, head coelomic cavity; ho, hood; lp, lateral plate; mb, bilobate muscle (musculus bicornis); mcoa, constrictor muscle of the mouth (musculus constrictor oris alter); mcop, primary constrictor muscle of the mouth (musculus constrictor oris primus); mdve, external vestibular dilator muscle (musculus dilator vestibuli externus); mep, multilayered head epidermis; mes, musculus expansus superior; mo, mouth opening; mos, mediolongitudinal-oblique head muscles (musculus obliquus superficialis); myf, myofilaments; nu, nucleus; opn, optic nerve; pep, perioral epidermis; syc, synaptic cleft; syv, synaptic vesicles.

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7.4 Histological structure, cytology, and functional significance of organ systems 

Rieger et  al. (2011) suggested a structural and functional subdivision of the chaetognath brain into two domains, one posterior brain domain that may be primarily involved in the integration of sensory input, and the anterior brain domain that may be involved in the control of the mouthparts and the anterior part of the digestive system. This claim is supported by developmental aspects of the cephalic nervous system (see Section 7.5.4, “Neurogenesis”). The circum­ esophageal arrangement of the adult cephalic nervous system, including, in addition to the brain, vestibular and subesophageal ganglion (compare Fig.  7.11), has already been recognized by Nielsen (2001). However, the situation in the hatchlings clearly shows that we do not only face one but two brain components that have a basically circumoral/ esophageal arrangement (Rieger et al. 2011).

7.4.3.2 Ventral nerve center The ventral nerve center (VNC, syn.: “ventral ganglion”) is found ventrally in the midregion or in the first third of the trunk depending on the species considered (e.g., Figs. 7.26 and 7.31 E). A VNC seems to be absent only in the bathypelagic species Bathybelos typhlops, along with some sense organs such as eyes and the corona ciliata. The central nervous system of B. typhlops is thus restricted to the elongated head and consists of the terminal brain (dorsal), the posterocephalic “dorsal ganglion”, and the circumesophageal chain of mediocephalic ganglia (Owre 1973; Bieri & Thuesen 1990). The invention of an additional dorsal ganglion and overall shift of components of the central nervous system to dorsal part of the body is considered an interesting convergence to hemichordates and chordates and most probably because of functional adaptation to low-oxygen environments (Bieri & Thuesen 1990).

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The VNC is connected to the brain via the paired, gently descending main connectives, which surround the posterior part of the esophagus as well as the anterior part of the midgut and finally enter the VNC at its anterolateral corners (e.g., Goto & Yoshida 1987; Harzsch & Müller 2007; Harzsch et al. 2009) (see Figs. 7.26, 7.29 A, 7.30  C, 7.31  C–D, and 7.39  B). At either side of the VNC, neurite bundles project into the intra- and basiepidermal neuronal plexus, which receives input from ciliary fence and tufts organ scattered in the whole epidermis (see Section 7.4.1 for details). This connection of basiepidermal plexus profiles and the likewise basiepidermal VNC makes it reasonable to assume that the VNC controls swimming by initiating contractions of the body wall musculature and coordinates mechanosensory input from the numerous ciliary fence and ciliary tuft receptors (reviews in Bone & Pulsford 1984; Goto & Yoshida 1984, 1987; Shinn 1997; Perez et al. 2014; Harzsch et al. 2016). In Spadellidae, two pairs of neurite bundles, called caudal nerves, project posteriorly from the VNC into the tail in bilaterally symmetrical fashion (Figs.  7.26  B, 7.30  C, and 7.32 E). To the posterior end of the tail, both pairs of caudal nerves bend medially and fuse, thus forming two, horizontally stacked caudal loops (Harzsch et al. 2009). Generally, the VNC shows three distinct macrocompartments: a fibrillar core neuropil and two lateral aggregations of neuronal somata flanking the core neuropil (Goto & Yoshida 1987; Bone & Goto 1991; Shinn 1997; Harzsch & Müller 2007; Harzsch et al. 2016) (Figs. 7.30 A–B and 7.31  F). Lateral aggregations of neuronal somata are also subcompartimentalized into numerous circular, drop-shaped, or columnar clusters stacked and aligned relative to the trunk’s longitudinal axis (e.g., Hertwig 1880; Goto & Yoshida 1987; Harzsch et al. 2009). For a long time, cellular constituents and specific ultrastructures in

◂ Fig. 7.29: Location and ultrastructure of the brain (br) and associated afferent nerves in the head of Spadella cephaloptera (A–B, D–G) and Parasagitta setosa (C). A, Mediosagittal semithin (LM) section of the head and trunk documenting the dorsal location of the brain and ventral position of the ventral nerve center (vnc). Individual has been fixed in relaxed condition; thus, hood (ho) is not retracted. B–G, Original TEM micrographs. B, Slightly oblique, parasagittal section of posterior (sensory) domain of the brain showing the core neuropil (cnp), clustered neuronal somata (nso), and a sensory cavity of retrocerebral organ (ro). Basiepidermal location of the brain is indicated by resting ventrally on the epidermal extracellular matrix (ecm), and dorsal coverage is achieved by extremely flattened proximal epidermal cells serving as sheath cells (fshc). C, Transverse section of posterior (sensory) domain of the brain with a cluster of neuronal somata in high-power magnification; somata a separated from each other by glial sheaths (gls) being radial projections of sheath cells (shc) located at periphery of the brain. D, Detail of core neuropil (region indicated by dotted rectangle in B); neuropil contains numerous synaptic profiles identified by the presence of axonal terminals (axt), including synaptic vesicles (syv), synaptic clefts (syc), and adjoined dendrites (den); larger vesicles with highly osmiophilic contents (nsv) reveal common presence of neurosecretory cells. E, Transverse section of most posterior part of the brain at the level of retrocerebral organ showing both sensory cavities filled with branched ciliary processes (ro), surrounded by somata of brain neurons and retrocerebral receptor cells; sensory cavities are flanked medially by coronal nerves (con) and laterally by optic nerves (opn, the right one only partly visible). F, Close-up of optic nerve in cross section. G, Close-up of a coronal nerve (indicated in overview section E by dotted rectangle). Further labels: cc, corona ciliata; es, esophagus; hco, cephalic coelomic cavity; int, intestine; mep, multilayered head epidermis; mi, mitochondrion/mitochondria; mo, mouth opening; mos, mediolongitudinal-oblique head muscles (musculus obliquus superficialis); ov, ovary; pec, proximal epidermal cells; ph, pharynx; trco, coelomic cavity of the trunk; tlmu, longitudinal (primary) trunk musculature.

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Fig. 7.30: Trunk nervous system of chaetognaths. A, B, Immunolocalization of synapsin (SYNORF 1; green) in a whole mount of the ventral nerve center of adult Parasagitta setosa combined with a nuclear marker (red; ventral views; A is black-white inverted). Synaptic contacts are confined to the core neuropil which shows a highly ordered subdivision into ca. 80 serially arranged microcompartments. C, Localization of RFamide-like immunoreactivity in the ventral nerve center of Spadella cephaloptera (ventral view of whole mount specimens, black-white inverted confocal laser-scan images). Identifiable neurons are encircled and labeled with letters. Arrows point to additional, unidentified neurons. Abbreviations: CO, main connective; CT, caudal tract (= caudal loop); IB, intermediate bundle; LB, lateral bundle; MB, medial bundle. Reprinted from Perez et al. (2014).

the VNC were just randomly described by using pelagic Sagittidae as study models (Bone & Pulsford 1984; Goto & Yoshida 1987; Bone & Goto 1991). Ahnelt (1980, 1984) and especially Harzsch et al. (2009) contributed insights from ultrastructure of S. cephaloptera. (Immuno)Histochemistry provided further details of the spatial distribution and interconnection of neuronal cell clusters within the VNC of various chaetognaths. Besides elegantly separating neuronal somata and neurites from each other, the

coherence and patterning of specific types of neurons could be documented by means of their neurotransmitter and neuropeptide equipment, according to neurophylogenetic methodologies outlined by Kutsch and Breidbach (1994) and refined later on by Harzsch (2006a, b) and Loesel and Richter (2014). For instance, disparity of RFamide-like immunoreactivity, among other markers, may bear remarkable potential to resolve chaetognath neurophylogeny (Harzsch et al. 2009, 2016; and section below).

7.4 Histological structure, cytology, and functional significance of organ systems 

Based on TEM analysis of fine-scaled, layer-cut ultrathin sections, Harzsch et  al. (2009) identified six different types of neurons in the VNC of S. cephaloptera. (1) Large neurons (“ln”) may be solitary or occur in small groups of up to three cells. The ln-neurons mostly occupy a position close to the ventromedian rim of the lateral clusters (Fig. 7.31 F). Their immediately branching neurites display a large diameter and reach deeply into the VNC neuropil where they take on a longitudinal course. This feature combined with the large size of these neurons, and their high metabolic activity, as indicated by high abundance of tubular mitochondria and Golgi stacks as well as a strongly developed rough endoplasmic reticulum reveal their probable function as motoneurons or command neurons. Based on soma shape, its position in the cluster, the nuclear structure (shape, eu-/ heterochromatin ratios), diversity of cytoplasmic organelles, and transverse projection patterns of neurites (from soma into neuropil), five further types of small neurons are distinguishable (“sn1–5”). (2) The “sn1” neurons are usually found at the lateral (facing the epidermal ECM) rim of the cluster; the nucleus contains minor portions of heterochromatin and thus appears rather inconspicuous in the cytoplasm (Figs.  7.31  F and 7.32  A, D). (3) The poorly organelle-supplied “sn2” neurons are encountered in small groups in medioventral position close by the core neuropil (Figs.  7.31  F and 7.32  A); in sagittal section, they appear aligned across clusters resembling a string-of-pearls formation. (4) The “sn3” neurons are restricted to the (inner) medial half of the lateral aggregations of neuronal somata; their cytoplasm contains tubular, often branched mitochondria; the nucleus displays considerable portion of heterochromatin. (5) The “sn4” neurons are commonly found medioventrally in places where the neurite bundles break through the glial sheath and feed into the core neuropil; the nucleus includes largest portion of heterochromatin among all VNC neurons described. (6) The “sn5” neurons widely resemble “sn4” neurons in shape and cytoplasmic composition but clearly differ from the latter by their position in each cluster (always adjoining and partly intertwining with the large motoneurons [“ln”] and intense glial wrapping along their entire path). The often drop-shaped or columnar lateral VNC neuron clusters are separated from each other by extremely flattened, microtubule-enriched radial (medial) processes of the basalmost proximal epidermal cells, thus acting as glial-like sheath cells (Fig. 7.32 A–B). These sheath cells also establish a septum between medial (inner) face of the lateral somata clusters of VNC-somata and the core neuropil (Fig.  7.32  C). This septum is pierced (“serially fenestrated”: Harzsch et al.

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2009) in places where neurite bundles project from the somata clusters (Figs. 7.31 F and 7.32 D). The core neuropil of the VNC is composed of highly ordered subdivision into ca. 80 serially arranged synaptic microcompartments (Harzsch & Müller 2007) (Fig.  7.30  A–B). Furthermore, immunolocalization of RFamide-related molecules provides evidence for the presence of serially arranged, individually identifiable neurons that are homologous among the various hitherto investigated species of Sagittidae (including P.  draco) and Spadellidae (Harzsch et  al. 2009), suggesting that the potential to generate serially arranged neurons with individual identities is part of the chaetognath ground pattern (Fig. 7.30 C). This view is supported by the finding of numerous septum-like protrusions of glial-like proximal epidermal cells (see previous paragraph and Harzsch et  al. 2009). In P. gotoi, Goto et  al. (1992, compare their fig. 1f) described a single RFamide-like immunoreactive bipolar neuron located in the tail region that connects to both caudal loops, but this observation has not been made in S. cephaloptera, which has a similarly unpaired fiber that innervates the tail region but with no trace of a neuronal cell soma (Harzsch et al. 2009). The VNC is a dominant organ in hatchlings and extends over half of the body length, whereas the brain is still rather rudimentary. Goto et  al. (1992) and Rieger et  al. (2011) have characterized the postembryonic development of RFamide-related molecules with immunofluorescence methods. Perez et al. (2013) have analyzed the proliferation of neuronal progenitor cells in the developing central nervous system (see Section 7.5.4, “Neurogenesis”).

7.4.3.3 Intra- and basiepidermal neuronal plexus The extensive basi- and intraepidermal neuronal plexus represents the peripheral nervous system of chaetognaths (Figs.  7.15  A and 7.33). The neuronal plexus displays two interconnected horizontal layers of aggregated neurons or neurites, (1) a basiepidermal domain with more or less orthogonally arranged neurite bundles (Figs. 7.15 A, 7.17 B, 7.18 A, and 7.33 B–D) and (2) an expanded intraepidermal domain (Figs. 7.15 A, 7.17 A, and 7.33 A). The first and more complex layer is found close by the thick and fibrillous epidermal ECM and comprises transverse, oblique, and longitudinal neurite bundles the connection pattern of which somewhat resembles an orthogon (Ahnelt 1984). Figure 7.33 C in this chapter illustrates a neurite bundle in horizontal section being an element of this basiepidermal orthogon. The second plexus domain is found crammed into the transition zone of distal and proximal epidermal

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7.4 Histological structure, cytology, and functional significance of organ systems 

cells without showing any kind of geometric arrangement (Figs. 7.17 A and 7.33 A). There, also neuronal somata are recognizable by their characteristic shape and cytoplasmic appearance (Shinn 1997) (Figs. 7.15 A and 7.33 A). The distal and proximal aggregation zone is interconnected by a highly complex network of plexus profiles (neurites or neurite bundles), which pass through the tiny interspaces of the proximal epidermal cells and are surrounded by extracellular filamentous material and can be comprehensively visualized using antitubulin immunohistochemistry (Harzsch & Müller 2007; compare also Fig. 7.15 A–B, F in this chapter). The intra- and basiepidermal neuronal plexus is particularly dense and noticeable in the vicinity of ciliary sense organs, as for instance, the ciliary fence and tuft receptors (Fig. 7.33 D). Although extensive intraepidermal plexus profiles are common in various metazoan phyla (review of Richter et al. 2010), the combination of a mainly orthogonal basiepidermal plexus and a heavily diversified intraepidermal plexus may be inevitably linked to a pluristratified (multilayered) epidermis. As thickness and structural complexity is uniquely high in the chaetognath multilayered epidermis, it would not be surprising if the basi- and intraepidermal neuronal plexus turned out to be highly derived, too. Consequently, the combination of a multilayered epidermis and two-level neuronal plexus may be interpreted as an additional apomorphy of this animal group. In Chaetognatha, acetylcholine is known to be the major neuromuscular transmitter that reaches constituting cells in the primary longitudinal musculature of the trunk

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and tail by diffusion through the epidermal ECM to elicit muscle contraction (Bone & Goto 1991; Shinn 1997). The cholinergic neurites are part of the profuse intra- and basiepidermal neuronal plexus described above (Duvert & Barets 1983; Bone & Pulsford 1984; Bone et  al. 1987b; Duvert & Salat 1990a; Bone & Goto 1991; Duvert et al. 1997; Shinn 1997; Harzsch & Müller 2007). This extensive intraand basiepidermal neuronal plexus both in the head region and in the trunk can also be visualized by antiaspartate immunohistochemistry (Duvert et  al. 1997). As mechanosensory input from the numerous ciliary fence and tuft receptors in the epidermis feeds into the plexus (Bone & Pulsford 1984; Bone & Goto 1991; Shinn 1997), it appears likely that the intra- and basiepidermal neuronal plexus acts as a functionally semiautonomous system that mediates the sensory-motor integration.

7.4.4 Muscular apparatus and locomotion In accordance with preferred lifestyle as ambush predators, the system of chaetognath locomotory muscles is complex and, as innervated by the likewise complex central nervous system (see Section 7.4.3, “Nervous System”), is capable of delivering fast movements. Duvert (1989) even considered this animal’s ability to react fast as ranking among the fastest invertebrates yet studied in this respect. Compared with other organ systems, the somatic musculature, comprising both locomotory (e.g., Figs. 7.34– 7.35 and 7.39 A, B) and visceral muscles (Figs. 7.34 D, 7.40 C,

◂ Fig. 7.31: Ultrastructural aspects of nervous system components found at posterior end of the head (A–B) as well as the trunk (C–F) in various chaetognath species: Ferosagitta hispida (A, C), Spadella cephaloptera (B, E–F), Parasagitta setosa (D). Originals. A–B: Stomatogastric nervous system. A, Median sector of semithin (LM) cross section of posterior cephalic region. B, Most anterior part of stomatogastric nerve projecting posteriorly from subesophageal ganglion, regular neurites (neu) and profiles of giant neurons (gn) are cut longitudinally and multiply enveloped by glial sheath cell processes (gls), sector shown exemplarily by dotted rectangle in A. TEM. C, Left half of semithin (LM) cross section of anterior head region showing basi-epidermal position of left main connective (mc); note the breakthrough in lateral region of cuticle (cu) marking position of vestibular pit (vep). D, Overview TEM cross section of main connective in anterior trunk region, connective is enwrapped by epidermal extracellular matrix (ecm), epidermis mostly ripped off due to fixation artifact. E, Semithin (LM) cross section of posterior trunk region containing ventral nerve center with two lateral clusters of vnc somata (vnso) and central vnc neuropil (vcnp). F, Overview TEM cross section of right half of ventral nerve center showing some types of vnc neurons: large neurons (ln) with a multisegmented nucleus (nu) and numerous mitochondria (mi) at ventromedian rim of the somata cluster, small type 1 neurons (sn1) with spherical nuclei containing high portions of heterochromatin, and small type 2 neurons (sn2) with polymorphous nuclei containing low portions of heterochromatin; multilayered glial sheath (gls) separate somata cluster from central neuropil and compartmentalize groups of neurons within the clusters, the glial sheath is continuous with most proximal, extremely flattened epidermal cells functioning as glial sheath cells (shc). Glial sheath has breakthroughs (arrowhead) where different sorts of neurite bundles project into vnc neuropil. Further labels: br, brain; cemu, circumesophageal muscles; csec, circumesophageal commissure; cfo, ciliary fence organ; dec, distal epidermal cells; es, esophagus; gs, grasping spines; hco, head coelomic cavity; hmu, head musculature; ho, hood; int, intestine; lp, lateral plate; mb, bilobate muscle (musculus bicornis); mcop, primary constrictor muscle of the mouth (musculus constrictor oris primus); mdve, external vestibular dilator muscle (musculus dilator vestibuli externus); mep, multilayered epidermis; mos, mediolongitudinal-oblique head muscles (musculus obliquus superficialis); pec, proximal epidermal cell(s); pep, perioral epidermis; stn, bundle of small transverse neurites; trco, trunk coelomic cavity; tlmu, primary longitudinal (trunk) musculature; vsg, vestibular ganglion (anterior end).

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7.4 Histological structure, cytology, and functional significance of organ systems 

and 7.41  B), has been investigated intensively across all chaetognath subtaxa, and among them especially in Sagittidae and Spadellidae (Hertwig 1880; Grassi 1883; Ritter-Záhony 1909; Burfield 1927; John 1933; Kuhl 1938; Hyman 1959; Duvert & Salat 1979, 1980; Duvert 1991; Bone & Duvert 1991; Duvert & Salat 1995; Shinn 1997; Casanova & Duvert 2002; Casanova et  al. 2003). The vast majority of chaetognath muscles are cross striated, but few smooth muscles were documented as well. Transmission electron microscopic examinations revealed many types of cross striation, indicating the way the sarcomeres are formed and arranged with respect to each other (Fig.  7.34  A–B). However, chaetognaths lack oblique-striated muscles, which are typical for many worm-like and soft-bodied invertebrates, such as nematodes, annelids, priapulids, or mollusks (for a review, see Schmidt-Rhaesa 2007). Supercontraction was documented by specialized cross-striated, transverse, and longitudinal muscles in some subtaxa of Phragmophora (see Casanova & Duvert 2002 and below).

7.4.4.1 Gut-associated muscles Visceral muscles enwrap the entire gut system. Visceral muscle fibers are cross striated and quite distinct around the esophagus, whereas they appear less obvious in the intestinal region (Figs.  7.34  D, 7.40  C, G, and 7.41  B, E). Circum­intestinal muscle cells show more myoepithelial characteristics. However, in contrast to subepidermal myoepithelial (coelothelial) cells, more and much denser packed myofilaments are encountered (compare Fig. 7.40 G with Fig. 7.40 H). Sphincters surrounding the anus as well as the female gonopores are made of myoepithelial cells (Duvert & Salat 1995). For further details, please see section further below on the gut system (see Fig. 7.37 A, F).

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7.4.4.2 Cephalic locomotory muscles Regarding locomotory muscles, comprehensive insights were provided by Casanova and Duvert (2002) who analyzed semithin histological sections as well as TEM data of not less than 47 species. Muscular patterns are highly complex and a multidirectional arrangement in the head feature. At least 16 head muscles are known. They maintain the shape and posture of the head and also move the grasping spines and teeth (e.g., Shinn 1997) (some relevant cephalic muscles are given on Figs.  7.3  A–B, 7.19  A, 7.21  B, and 7.28  A, D). “Myotendinous junctions” (complexus lateralis sensu Duvert & Barets 1983) attach the cross-striated myocytes to the epidermal ECM in various places of the head suggesting these cells to derive from coelothelial cells of the embryonic head coelom (Hertwig 1880; Doncaster 1902; John 1933; Shinn 1997, the developing head mesoderm is illustrated in fig. 10.10 of Harzsch et al. 2015). Contraction of head muscles is initiated either by regular neuromuscular synapses formed by motor neurons, which are mainly accommodated in the vestibular ganglia and projecting via the dorsal (Fig.  7.28  E) and mandibular nerves (e.g., Goto & Yoshida 1987; Shinn 1997) or by the various axon profiles of the intra-/basiepidermal nerve plexus (e.g., Fig. 7.33). In the latter case, some myocytes, such as those of the unpaired bicorneal muscle (musculus bicornis sensu Ritter-Záhony 1909; see also Fig. 7.21 B in this chapter), protrude through lobular invaginations of the epidermal ECM and establish close synapses with plexus motor neurons. Neurotransmitters have then to diffuse through a locally thinned epidermal ECM (Duvert & Barets 1983; Shinn 1997). More details on neuromuscular innervation can be found in section above on nervous systems.

◂ Fig. 7.32: Ultrastructural aspects of ventral nerve center (A–D) and caudal nerves (E) in the trunk and tail of Spadella cephaloptera. Original TEM micrographs. A, Slightly oblique parasagittal section of ventral nerve center spanning a cutting range from transition zone of lateral somata cluster (vnso) and core neuropil (vcnp) on the left side toward mediosagittal plane on the right. Septa (gls) built by glial sheath cells (shc) leave openings through which neurite bundles of small (stn) and large diameters project transversally into the core neuropil, glial sheaths also subgroups neuronal somata within the cluster, one subcluster is marked by arrowheads. B, Close-up of ventral periphery of core neuropil; vnc neurites are multiply wrapped by radial projections (gls) of peripheral sheath cells (shc); note the difference in electrondensity of tonofilament-rich cytoplasm of sheath cells in comparison with syntypic proximal epidermal cells (pec). C, Transition zone of core neuropil (on the right) and somata cluster (on the left) showing soma of an internalized sheath cell in high-power magnification. D, Close-up tangential view of immediate transition zone between core neuropil and lateral cluster of vnc somata; glial sheath septa are ramified and leave openings for transverse neurite bundles (e.g., stn, bundles of small transverse neurites) to pass through and feed into the core neuropil; sector exemplarily shown in A by dotted rectangle. E, Cross section of lateral periphery of midtail region showing base of lateral fin (lf) and lateral aspect of tail coloemic cavity (taco), including some aggregates of mature sperm (spa) or various spermatogenic stages (spta); caudal nerve (cdn) is split in two branches dorsal and ventral of the fin core (fc) base. Further labels: dec, distal epidermal cells; ecm, extracellular matrix (of the epidermis); lfc, lateral field cells; ltn, bundle of large transverse neurites; nu, nucleus; pec, proximal epidermal cells; rc, ray cell; sn1, small type 1 vnc neuron; sn2, small type 2 vnc neuron; sn3, small type 3 vnc neuron; tlmu, primary longitudinal (trunk) musculature.

234 

 7 Chaetognatha

A

B dec

tfa

ipne

sg ipso

pec shc

mi

v

ecm

axt

nu

bpp

go

syv

pec

nu

hemu

1 µm

ipne

0,5 µm

D

C

ci

ecm

pec cfo

pec

ecm

edr

bpnb

bpnb pec

pmyc

2 µm

hemu

2 µm

Fig. 7.33: Ultrastructure of main components of neuronal intra- and basiepidermal neuronal plexus of two chaetognath species (A: Spadella cephaloptera; B–D: Ferosagitta hispida) as depicted from TEM. A, Intraepidermal plexus domain: cross section of most distal part of dorsolateral tail epidermis exhibiting a soma of an intraepidermal plexus neuron (ipso) tapering into small neuritic processes at either side (ipne), plexus neuron is located between distal epidermal cell unilayer (dec) and distalmost layer of proximal epidermal cells (pec). B–D: Basiepidermal plexus domain. B, Cross section of basiepidermal neurite bundle in dorsolateral trunk region, encapsulated by glial sheath cell (shc) and sitting on the epidermal extracellular matrix (ecm), neurite bundle includes an axonal terminal (axt). C, Obliquetangential section of most basal part of proximal epidermal cells (in dorsolateral trunk region), slightly above the ecm, providing horizontal view of a basiepidermal plexus neurite bundle (bpnb). D, Basiepidermal plexus neurite bundle of thicker dimension (as compared with C) immediately below a ciliary fence organ (cfo). Further labels: ci, receptoral cilia; edr, extremely electron-dense distal root body; go, Golgi stack; hemu, paramedian, heterosarcomeric, secondary tail musculature; mi, mitochondrion; nu, nucleus; pmyc, mesodermal perimysial cells; sg, secretory granule; syv, synaptic vesicles; tfa, tonofilaments; v, vacuole.

7.4 Histological structure, cytology, and functional significance of organ systems 

7.4.4.3 Trunk and tail locomotory muscles Most locomotory muscles are arranged longitudinally in the trunk (e.g., Figs. 7.4, 7.31 E, 7.34 C, and 7.42 A) and tail (e.g., Fig.  7.35  A, 7.39  A, and 7.42  B), but some transverse muscles do also exist in these parts of the chaeto­ gnath body (see Figs. 7.4 B, 7.42 A, and description below). According to their course and topographical location relative to each other, locomotory trunk and tail muscles can be classified as primary or secondary muscles (already noticed by Grassi 1883). The highly specialized transverse or oblique muscles represent further type of locomotory muscles present in the trunk and tail regions. 7.4.4.3.1 Primary muscles Primary muscles represent the predominant tissues in the trunk and tail, by occupying approximately 80% of the entire volume (Duvert 1989; Shinn 1997). They are arranged in two, widely continuous (if at all only “interrupted” by the trunk-tail septum) pairs of muscular bands (“muscular quadrants” sensu Casanova & Duvert 2002). A dorsolateral pair is subjacent to the dorsal hemisphere of the integument, whereas the slightly thicker pair of ventrolateral primary muscles is located below the gut and delimits the coelomic cavities bands (Figs. 7.4 A, 7.34 C, 7.35 A, 7.39 A, and 7.42 A–B). Therefore, the primary muscles are a major component of the somatopleura, namely, the subepidermal coelothelial cells opposing the gut. Further coelothelial cells with myoepithelial characteristics adjoining the primary muscles along the dorsal and ventral midline are the medial myoepithelial cells (Fig. 7.39 B). Myoepithelial lateral field cells border these muscles at either side of the body (Shinn 1997) (Fig. 7.17 C–E). Primary myocytes display classic cross striation thus comprising alternating bands along the body’s longitudinal axis and are composed of a single type of sarcomeres (ultrastructural details on myofilaments can be depicted from the study of Duvert & Savineau 1986). However, the arrangement of sarcomeres, the distribution of metabolic compartments in the myoplasm, and the arrangement of sarcomeres (rela­ tive to endomembrane system) may be different. Hence, two different types of myocytes (type A and type B myocytes) are distinguished in the primary musculature of all chaeto­gnaths (Shinn 1997; see Fig. 7.34 B in this chapter), except for Spadellidae, which only show primary muscles made of type A myocytes (Duvert & Casanova 1994; Casanova & Duvert 2002). Type A myocytes (Figs. 7.34 B, E–F and 7.35  B–C), termed also A-fibers (Bone & Duvert 1991; Shinn 1997), are equally sized and display mitochondria that are restricted to peripheral areas and aggregate around the single nucleus (found apically immediately

 235

below adjoined coelothelial cells), their strap-shaped myofibrils are arranged in parallel. By contrast, type B myocytes (called B-fibers by Bone & Duvert 1991; Shinn 1997) are interspersed among groups of type A myocytes. Type B myocytes usually vary in size; their myoplasm comprises pockets rich in mitochondria and other organelles nested between the myofibrils (not restricted to the periphery) (Fig.  7.34  B, G–H). In addition, the sarcoplasmic reticulum shows higher complexity (see fig. 4.4. in Bone & Duvert 1991 for comparison of type A and type B myocytes, and Fig. 7.34 B of this section). In P. setosa, only 15% of primary myocytes belong to type B class. Their portion, however, may increase in posterior regions. In the same species, two groups of type A myocytes alternate with each group of type B myocytes in the trunk, whereas in the tail, just a single group of type A myocytes follows a single group of type B myocytes. Further variations in type A and type B ratio in primary musculature are related to the ecology of chaetognaths. Generally, type B myocytes were observed to be more voluminous in planktonic taxa, such as Sagittidae, than in benthoplanktonic ones (Duvert & Savineau 1986; Duvert & Casanova 1994; Shinn 1997; Casanova & Duvert 2002). Primary myocytes and among them the type A myocytes, in particular, are tied firmly to each other by a set of adhering structures that may include regular belt desmosomes (zonulae adhaerentes), gap junctions, as well as macular and zonular columnar junctions. Both types of columnar junctions are characterized by peculiar intercellular domains revealing a system of repetitive, fibrillous “columns”. These seam-like columnar junctions are unique in the animal kingdom. In type B myocytes, however, only gap junctions and macular columnar junctions can be found (Duvert et  al. 1980) (Fig.  7.34  G). The primary musculature as a whole is capable of delivering rapid but brief contractions (Goto & Yoshida 1981). Thus, the primary ­musculature is considered to be mainly responsible for directional jerking-swimming movements of chaetognaths (e.g., Bone & Duvert 1991). It is assumed that differential, most likely alternating contractions of dorsal and ventral primary muscles in particular quadrants of the trunk and tail induce rhythmic bursts causing the peculiar way the chaetognaths move (Bone & Duvert 1991). Type B myocytes with typically higher abundance of mitochondria are assumed to enable sustained contractions and are consequently suitable to sustained jerking-swimming activities in a pelagic environment (Duvert 1989; Casanova & Duvert 2002). The fact that type B myocytes are absent in representatives of benthic taxa like Spadella and Paraspadella with generally reduced swimming activities may be taken as an ecological trait supporting this assumption.

236 

A

 7 Chaetognatha

B

1

2: A-fibers

1: B-fibers

2 3 4 5

C

tlmu

tlmu

mep

D

pis

gec ov

int

trco

E

tlmc

ov

lf

tlmu

D

tlmu

gmu

1 µm

100 µm

myf

F

sr

myf

A-b A-b

sr

H-b mi

I-b 1 µm

G

0,5 µm

cmj sr

1 µm

H mi

myf

iv

mi

myf

1 µm

7.4 Histological structure, cytology, and functional significance of organ systems 

7.4.4.3.2 Secondary muscles The rather inconspicuous secondary musculature constitutes only 1% of the trunk’s and tail’s tissues as observed in P. setosa and P. friderici by Duvert (1991). Three pairs of secondary muscle bands are found in bilaterally symmetrical formation: (1) the dorsomedian pair is crammed into the corner space at either side of most dorsal aspect of the dorsal mesentery, the dorsomedian part of the epidermal ECM (faces these muscles dorsally), and the dorsal primary muscle bands (face these muscles ventrally and laterally) (in various representatives of Phragmophora with more dorsolateral position slightly distanced from mesentery); (2) the ventromedian pair mirrors the former configuration by being adjoined by respective structures in the ventral hemisphere (except representatives of Phragmophora in which ventral secondary muscles take a ventrolateral position and are also lined by ventrolateral end of the transverse muscles); and (3) each one secondary muscle of the lateral pair is seen at the dorsolateral corner of the ventral hemisphere lined by ventrolateral primary muscle and coelothelial cells of the lateral field (compare fig. 1 in Duvert 1991 or fig. 1 in Casanova & Duvert 2002). The position of secondary muscle bands in trunk and tail of S. cephaloptera is exemplarily indicated in Figure 7.42 B. Myocytes of secondary muscles in general are cross striated and show an enormous disparity in ultrastructural organization and arrangement of sarcomeres (compare Fig. 7.34 A1–4). They become differentiated rather late in development; the sarcomeric pattern appears not earlier than 2–3 days after hatching, thus being delayed in relation to primary muscles (Casanova & Duvert 2002). Three types of crossstriated myocytes are known (for a review, see Casanova & Duvert 2002): (1) classic heterosarcomeric myocytes (Fig.  7.34  A1), (2) hybrid or irregularly heterosarcomeric

 237

myocytes (Fig.  7.34  A2), and (3) homosarcomeric myocytes (Fig.  7.34  A3–5). Homosarcomeric sarcomeres may occur in secondary muscles of few representatives of Heterokrohniidae (X. sorbei) and some genera of Spadellidae (Paraspadella spp. and Spadella spp., Figs. 7.34 A4 and 7.35  D). These sarcomeres contain discontinuous Z-discs and are considered pleomorphic; thick filaments or groups of thick and thin filaments may invade the aligned neighbor sarcomeres, leading to an even tighter constriction of A-bands. Hence, homosarcomeric secondary muscles are considered a “supercontracting machinery”. By contrast, two different types of sarcomeres (S1- and S2-sarcomeres) are aligned in heterosarcomeric muscles (Figs.  7.34  A1 and 7.35  B–C). They were documented frequently in representatives of the Aphragmophora, including Krohnittidae (Krohnitta spp.) and most likely all Sagittidae (e.g., Sagitta spp., Ferosagitta spp., Flaccisagitta spp., Parasagitta spp., Aidanosagitta spp., and P. draco). Moreover, but less widespread on the species level, heterosarcomeric myocytes/muscles were also found in phragmophoran subtaxa, such as Heterokrohniidae (Heterokrohnia spp.), Eukrohniidae (Eukrohnia spp.), and Spadellidae (Hemispadella spp.). Heterosarcomeres are generally longer than those establishing the primary muscles (compare fig. 8 in Casanova & Duvert 2002). S1-sarcomeres show ultrastructural features typical of slowly contracting cross-striated myofibers as encountered in many invertebrates (Duvert 1991; Casanova & Duvert 2002; Royuela et al. 2003). Close resemblance not only applies to the remarkable length of the sarcomeres (Fig. 7.35 B–C) but also to the specific pattern of 9–10 thin myofilaments surrounding a single thick filament (Fig. 7.35 E). By contrast, S2-sarcomeres indeed possess thicker (10–15 nm) and thinner (ca. 6 nm) myofilaments but are unusually

◂ Fig. 7.34: Pattern, typology, and ultrastructure of cross-striated locomotory muscles (A–C, E–H) and visceral muscles (D). Light microscopy (LM): C; TEM: D–H. A, Schematic overview of sarcomeric ultrastructure and arrangement in cross-striated locomotory muscles of various chaetognaths: (1) classic heterosarcomeric, secondary muscle with aberrant S2-sarcomeres alternating with regular S1-sarcomeres, e.g., Sagittidae; (2) irregular heterosarcomeric, secondary muscle of Archeterokrohnia rubra with S1- and S2-sarcomeres alternating in irregular fashion; (3) classic homosarcomeric, transverse muscle (only S1-sarcomeres present), e.g., Spadellidae; (4) supercontracting homosarcomeric, secondary muscle: Spadella cephaloptera; (5) primary musculature with comparatively short S1-sarcomeres: all Chaetognatha. Adapted from Casanova and Duvert (2002). B, Schematic reconstruction of arrangement of sarcomeres and sarcoplasmic compartments in primary trunk and tail muscles of chaetognaths: (1) B-fibers with myofibrils (myf) alternating with sarcoplasmic spaces mostly occupied by mitochondria (mi), sarcoplasmic reticulum (sr) show longitudinal interconnections (anastomoses); (2) A-fibers contain more adjacent myofibrils, sarcoplasmic reticulum and including mitochondria are widely restricted to the periphery of the muscle cell, sarcoplasmic reticulum lacks anastomoses. Arrows mark entry pores into invaginated and expanded system of transverse tubules. Adapted from Bone and Duvert (1991), reprinted from Shinn (1997). C, Semithin cross section of posterior trunk region of Ferosagitta hispida. D, Medioventral periphery of the intestine (int, indicated by dotted square in C) showing bases of midgut epithelial cells (gec) and cross-cut profiles of surrounding peri-intestinal sinus (pis) and cross-striated visceral muscle (gmu). F. hispida. E–F, Detail of A-fibers of primary musculature in longitudinal (E: S. cephaloptera) and transverse (F: F. hispida) section; note the compression of H-band due to massive contraction of locomotory muscles during fixation. G, Close-up of B-fibers in cross section. Thick and thin myofilaments are strictly aligned. Myocytes are firmly attached via columnar macular junctions (cmj). F. hispida. H, Cross-cut B-fibers with high portion of mitochondria loosening up pattern of myofibrils. F. hispida. Further labels: A-b, A-band; I-b, I-band; iv, invaginations (transverse tubuli); lf, lateral fin; mep, multilayered trunk epidermis; ov, ovary; pm, plasma membrane (of muscle cell); trco, coelomic space of the trunk; tlmc, primary longitudinal (trunk) muscle cells, tlmu, primary longitudinal (trunk) musculature.

238 

 7 Chaetognatha

arranged in variable, but often parallel or tubular arrays (Fig. 7.35 F) and do not display A- and I-bands (Fig. 7.35 C). The definite molecular composition of these filaments is still unknown, but they were suggested to be cytoskeletal filaments with contractile properties (Duvert 1991). Indeed, various muscle proteins including actin (possibly corresponding to the thin filaments) were detected in S2-sarcomeres by immunohistochemistry (Royuela et  al. 2003), but there is no evidence yet for myosin or paramyosin, suggesting that the thicker myofilaments may represent a type of intermediate filament. Hence, Royuela et al. (2003) suggested that the S2 sarcomeres are not true muscular structures but rather represent “pseudosarcomeres” of unknown physiological role. Heterosarcomeric myocytes are by all means unique in the animal kingdom and can, therefore, be added to the extensive list of autapomorphies defining the Chaetognatha (e.g., Duvert 1991; Perez et al. 2014). Ultrastructural features (Z-discs, length, and arrangement of myofilaments) indicate that heterosarcomeric (secondary) muscles are slow reacting but should work similar to supercontracting muscles (Duvert 1991; Duvert & Casanova 2002). An interesting subtype of heterosarcomeric (secondary) myocytes is realized in secondary muscles in a subtaxon of the Heterokrohniidae, the genus Archeterokrohnia. Casanova and Duvert (2002) described considerable variation in the pattern of S1- and S2-sarcomeres in the species A. rubra. (1) There are true heterosarcomeric myofibrils with both S1- and S2-sarcomeres, but exhibit a different mode of alternation by having two contiguous S1-stacks followed by a very long unit of S2-sarcomeres (Fig.  7.34  A2). (2) However, some of the S1-sarcomeres may be nested within S2-sarcomeres causing an S1/ S2-hybrid formation. (3) S2-sarcomeres may be absent resulting in a classic S1-based, cross-striated formation.

S2-sarcomeres are present in places where the Z-discs between two adhering S1-sarcomeres are separated. However, other S1/S2-hybrid formations with straight and complete Z-discs also suggest that S1-sarcomeres as a whole may derive from compact S2-sarcomeres by transformation. If this was true, the irregular S2-sarcomeres would be an intermediate evolutionary stage or developmental precursor of the regular S1-sarcomeres (Casanova & Duvert 2002). With the powerful, sustained contractions of secondary muscles, an antagonist is formed that helps to keep and adjust the tension in the hydroskeleton, namely the inner pressure applied by coelomic fluids (cf. Casanova & Duvert 2002). In benthic and benthoplanktonic Phragmophora (Spadellidae, Heterokrohniidae), the secondary muscles, most probably in cooperation with the transverse muscles, are proposed to enable the chaetognath to lift up the body from the substrate (Casanova & Duvert 2002). The same authors suggested that the extremely heterogeneous ultrastructural appearance of the secondary muscles of A. rubra may reflect the plesiomorphic state present already in the last common ancestor of Chaetognatha. By gradual, function-induced loss of structural complexity, the homosarcomeric system in Spadellidae as well as the homogenously heterosarcomeric formation in Sagittidae and further groups of Aphragmophora may be explained. In the genus Paraspadella, lateral secondary muscles were most likely transformed in a gradual process leading to the evolution of appendicular muscles used in derived species (e.g., P. gotoi) to in order to erect the body by moving large and bilobed posteroventral appendages (for further details, see Section 7.4.1, “Epidermis”, and Casanova et al. 1995, 2003). In Xenokrohnia, the secondary muscles display a hybrid architecture of those subtypes described for the

▸ Fig. 7.35: Pattern and ultrastructure of secondary locomotory muscles in chaetognaths, seen from longitudinal (B–D) and transverse (E–F) section planes. A, Transverse semithin (LM) section of anterior tail region showing two coelomic cavities completely occupied by spermiogenic cell clusters, vacuolated epidermal cells (vec) are visible around dorsal midline, the ventral surface is riddled with ventral gland papillae (vgp). Spadella cephaloptera. B, Slightly oblique, parasagittal section very close to the mediosagittal plane showing classic cross-striated, longitudinal, primary (tail) musculature (tlmu) and paramedian, heterosarcomeric, secondary tail musculature (hemu) in direct comparison. Ferosagitta hispida. C, Close-up of heterosarcomeric (secondary) muscle (same muscle band as shown in B); regular, more osmiophilic S1-sarcomeres (S1) alternate with irregular, lower osmiophilic S2-sarcomeres (S2) having a different set of myofilament strands; presence of adjacent axonal terminals (axt) indicates innervations of secondary muscle by basiepidermal neuronal plexus domain; note the difference in size and accuracy of arrangement of S1-sarcomeres of secondary muscle and sarcomeric bands in homosarcomeric primary muscle. F. hispida. D, Homosarcomeric, secondary muscle in S. cephaloptera, muscle has same position as described in B, epidermis (to the right) contains section of type 2 receptor cells (rc2) of a ciliary fence organ. E, Ventromedian region of anterior tail epidermis and immediately subjacent tissues including perimysial myoepithelial cells (pmyc) in the center flanked by sections of paramedian, heterosarcomeric, secondary muscle cells (hemu) in cross section. Section is exemplarily indicated by dotted rectangle in A. F. hispida. F, Detail of the same cross-cut heterosarcomeric, secondary muscle as indicated in E; thick myofilaments in S1-sarcomeres exhibit a lattice-like arrangement regular for cross-striated myofibrils, whereas in S2-sarcomeres thick filaments are more aligned forming a pattern of parallel sheets and tubes (arrow). F. hispida. Further labels: dec, distal epidermal cells; ecm, extracellular matrix (of the epidermis); fc, fin core matrix; gls, glial sheath (built by shc); lf, lateral fin; mecm, median (mesenterial) extracellular matrix; mep, multilayered tail epidermis; mi, mitochondrion; shc, sheathing proximal epidermal cells; tame, tail mesentery; te, testis (spermatogenic aggregates in both coelomic cavities of the tail).

7.4 Histological structure, cytology, and functional significance of organ systems 

A

B

vec tlmu

tlmu tame

te

fc

tlmu

mep

te

tlmu lf 2 µm

50 µm

vgp

C

hemu

pec

tlmu

E

ecm

dec

 239

D

hemu

ecm

ecm homu

mecm S1 pmyc

axt S2 mi pec

rc2

tlmu shc

gls

2 µm

E

2 µm

F

hemu

S1 sr

pmyc pec

S2 mi

hemu

ecm

2 µm

0,5 µm

240 

 7 Chaetognatha

genera Sagitta versus Spadella. It was suggested that chaetognaths have explored their own evolutionary pathways in generating muscle diversity (Casanova & Duvert 2002) so that muscle architecture at the moment is likely not useful for specifically relating Chaetognatha to other metazoan phyla (Perez et al. 2014). We agree with Duvert (1991) that the ultrastructural features of muscle architecture at the moment do not help us to specifically relate Chaetognatha to other metazoan phyla, but we nevertheless can conclude that this taxon seems to have explored its own evolutionary pathways in generating muscle diversity. It is linked to the ventral nerve center, but mechanosensory input from the numerous ciliary fence receptors in the epidermis also feeds into the plexus (Bone & Pulsford 1984; Bone & Goto 1991; Shinn 1997), which seems to act as a functionally semiautonomous system that mediates sensory-motor integration.

7.4.4.3.3 Transverse muscles Transverse muscles are also called “phragms”. They occur in bilaterally symmetrical formation and stretch as thin, more or less extended bands across the coelomic cavities (Shinn 1997). The transverse muscles anchor into the epidermal ECM at dorsal rim of the lateral fields and at the medial face of ventral primary to the root of the ventral mesentery (longitudinal) muscular quadrants (John 1933; Bone & Duvert 1991; Shinn 1997; Casanova & Duvert 2002) (Figs. 7.4 B and 7.42 A). In many Heterokrohniidae (Archeterokrohnia spp., Heterokrohnia spp., Xenokrohnia spp.), two pairs of transverse muscles are present, one pair is located in the trunk, the other one is restricted to the anterior part of the tail (Shinn 1997). Therefore, Hetero­ krohniidae have also been termed Biphragmophora (Casanova 1985a). Eukrohniidae and Spadellidae only exhibit one pair of transverse muscles (Phragmophora: e.g., Tokioka 1965a, b; Gasmi et  al. 2014), either limited to the anterior trunk region (Eukrohnia spp.) or extending throughout almost the entire trunk (Spadella spp.: see Fig. 7.42 A in this chapter, Paraspadella spp., etc.). Pelagic Sagittidae (including P. draco) are devoid of transverse muscles (Aphragmophora: e.g., Tokioka 1965a, b). Transverse muscles are always cross striated (homosarcomeric) across some chaetognath subtaxa such as Eukrohniidae and Heterokrohniidae. The length of the homosarcomeres varies considerably between 2.3 µm (Heterokrohnia spp., X. sorbei) and 4.5 µm (A. rubra, Eukrohnia spp.) (Casanova & Duvert 2002). Sarcomeres in transverse muscles of Spadellidae have perforated Z-discs allowing for the thick myofilaments to cross them, thus causing supercontraction (Casanova & Duvert 2002) (Fig.  7.34  A4).

Different functions of transverse phragma muscles have been discussed, among them the support to perform crawling motions (Tokioka 1965a), to enable more sophisticated modes of swimming (Salvini-Plawen 1986) or to maintain rigidity of the entire body once raised in part or even flip into fully vertical position (Tokioka 1965a), as for instance, by raising appendicular muscles and ventral adhesive organs in species of Paraspadella.

7.4.4.3.4 Oblique muscles Shinn (1989, 1997) reported obliquely projecting transverse muscle bands in the posterior part of the trunk in F.  hispida. They generally have a similar course and attachment sites as found in transverse muscles of phragmophoran taxa, but they differ from the latter by including smooth myocytes and being detached from the coelomic borders of the ventral primary musculature (surrounded by own subepidermal branches of the epidermal ECM). These muscles are proposed to facilitate egg laying.

7.4.4.4 Neuromuscular systems Chaetognaths also show a large variation in the ultra­ structure of neuromuscular junctions that was suggested to match the diversity seen, e.g., in arthropods (Duvert & Barets 1983). At least three types of neuromuscular synapses have been described so far, two related to muscles in the head and one related to the trunk muscles. In the head, nerves emerging from the brain are embedded in supporting structures, these being either ramifications of the epidermal EMC or sheets of extremely flattened, gliallike proximal epidermal/sheath cells (see Section 7.4.1, “Epidermis”, and Section 7.4.3, “Nervous System”). These nerves, which target the esophageal and somatic (locomotory) head muscles, have standard nerve endings and neuromuscular junctions that display ultrastructural features similar to classic motor end plates (Duvert & Barets 1983; Shinn 1997). Therefore, the first type of synapses is related to the esophageal muscles, which are innervated by more or less classical neuromuscular contacts from extensions of nerves penetrating the muscle tissue (Duvert & Barets 1983). The second type of synapses is related to other muscles in the head, where extensions of muscle cells protrude through the epidermal ECM and extend toward axons to form synaptic contacts with them. By contrast, the primary (longitudinal) muscles of the trunk (see above) are completely separated from the epidermis and the nervous system by a thickened epidermal ECM, which is crossed neither by extensions of muscle

7.4 Histological structure, cytology, and functional significance of organ systems 

fibers nor by neurites (Figs. 7.17 B, 7.18 A, and 7.33 B). This means that in this third type of synaptic terminals, the axons do not have direct synapses with underlying locomotory muscle fibers, but that the presynapses are separated from underlying muscles by this thick, fibrillar ECM which was also termed “connective stratum” by Duvert and Barets (1983). This very specialized type of neuromuscular innervation is mirrored in the arrangement of acetylcholinergic fibers, which deliver their transmitter by axonal (presynaptic) terminals. These axons terminate on the epidermal side of the ECM and therefore diffusely bath the muscle cells with acetylcholine (Bone et  al. 1987b, Duvert et al. 1997). In Chaetognatha, acetylcholine is known to be the major neuromuscular transmitter, and it is obvious now that it reaches myocytes of the primary trunk musculature by diffusion through the epidermal ECM to elicit contraction (Bone & Goto 1991; Shinn 1997). As mentioned above, the cholinergic neurites are part of a complex intra- and basiepidermal neuronal plexus (Duvert & Barets 1983; Bone & Pulsford 1984; Harzsch & Müller 2007), which is linked to the ventral nerve center and functions as a semiautonomous system that mediates sensory-motor integration (Bone & Pulsford 1984; Bone & Goto 1991; Shinn 1997). Duvert and Barets (1983) discussed whether this unusual type of neuromuscular innervations of the chaetognath primary trunk musculature may have structural counterparts in other animals and evaluate evidence from Pogonophora and Annelida. However, more ultrastructural data are necessary to draw a meaningful conclusion on this issue, but such comparisons for the future may hold some potential for phylogenetic insights. At the moment, we conclude that together with the secondary musculature and the other unusual morphological, embryological, and genomic features, the uncommon third type of chaetognath neuromuscular contact may be taken as evidence for the long evolutionary distance that separates the Chaetognatha from its closest (unknown) metazoan relative.

7.4.4.5 Considerations about muscle evolution within the Chaetognatha Given the enormous disparity in the ultrastructure and pattern of sarcomeres, displayed by secondary muscles in particular, Casanova and Duvert (2002) advocated for a revision of the classification formerly introduced by Grassi (1883) and used later on by many other authors. According to the newly proposed nomenclature, the term secondary muscles should only be used

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in a topographic context, thus referring to bilaterally symmetrical, cross-striated muscles extending longitudinally through trunk and/or tail and arranged basolaterally of the primary muscular quadrants. Instead, the consequent usage of homosarcomeric and hetero­ sarcomeric muscles was recommended. The ability to perform supercontraction poses a further level of typological diversification in cross-striated muscles of chaeto­gnaths. Many invertebrate taxa comprise several types of myocytes and, in particular, many subtypes of cross striated ones (see compilation by SchmidtRhaesa 2007). It has therefore been concluded that in some groups, such as Gastrotricha having both crossand oblique-striated muscles, the evolution of striated muscle types is better explained by functional rather than phylogenetic constraints (Schmidt-Rhaesa 2007). However, heterosarcomeric muscles/myocytes are most likely a unique (apomorphic) feature of the Chaeto­ gnatha. Indeed, Casanova and Duvert (2002) could be correct in assuming that the hybrid heterosarcomeric myocytes of A. rubra may reflect the plesiomorphic state for Chaetognatha and mark the evolutionary precursor structure of regularly cross-striated muscles. Casanova and Duvert (2002) remained indecisive as to how the homo- and heterosarcomeric secondary muscle bands are built in detail. Either way, a specific study on muscle differentiation in Heterokrohniidae in general and A. rubra in particular is needed. More precisely, it would be beneficial to explore in a further study the postembryonic development of A. rubra, although hard to sample in the field, with the aim to analyze if S2-sarcomeres are newly integrated into S1-sarcomeres or substituted by transforming into S1-sarcomeres. Furthermore, it needs to be evaluated if the possession of supercontracting cross-striated (transverse) muscles may reflect an ancestral character state from which oblique-striated supercontracting muscles derived. So far, supercontracting cross-striated muscles or structural intermediates between cross and oblique striation have been found only in very few taxa among Arthropoda (e.g., Hexapoda: Osborne 1967; Tardigrada: Dewel et al. 1993; Walz 1974, 1975). However, Arthropoda are quite derived ecdysozoans. If the Chaetognatha were basal offshoots of protostomes as proposed recently by some authors (e.g., Nielsen 2012; Perez et al. 2014; this chapter), crossstriated myocytes with supercontracting capacities may be the evolutionary precursor of oblique-striated muscles found in numerous lophotrochozoan subtaxa (e.g., Nemertini, Mollusca, Annelida) and less often in some taxa of Ecdysozoa (e.g., Nematoda) and Deuterostomia (Crinoida) (see Tab. 5.1 in Schmidt-Rhaesa 2007).

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7.4.5 Digestive system Major contributions to the histology of the digestive system (Figs. 7.36–7.39) have been provided by John (1933) on S. cephaloptera and Parry (1944) on both S. cephaloptera and P. setosa. Dallot (1970) described the gut anatomy of about 40 species with special attention to the occurrence of vacuolated intestinal cells and structure of the intestinal diverticula and discussed the relevance of these features in the debate regarding the affinities within the phylum. Using light and electron microscopy, specific studies on the intestinal epithelium were devoted to different aspects: development of digestive processes and formation of fecal pellets in F. hispida (Reeve et al. 1975), cytomorphology of the intestine in P. elegans (Welsch & Storch 1983b) and F. hispida (Shinn 1997), enzyme histochemistry of the intestine in P. elegans (Welsch & Storch 1983b), buoyancy adaptation in P. elegans (Bone et  al. 1987a), cell junctions in P. setosa (Duvert et  al. 1980; Duvert & Gros 1982), visceral musculature in P. friderici and P. setosa (Duvert & Salat 1995), feeding and starvation experiments in S. cephaloptera (Perez 2000; Perez et  al. 2000), and degrees of vacuolation of the absorptive intestinal cells in S. bipunctata, Parasagitta megalophthalma, M. minima, and S. zetesios (Perez 2000; Perez et al. 2001). Ultrastructural observations of the whole digestive tract including the oral cavity and the esophagus have only addressed S. cephaloptera (Arnaud et al. 1996), P. setosa, S. serratodentata, and S. pacifica (Perez et al. 1999; Perez 2000). The digestive tract of chaetognaths is divided into two functional units in respect to the distribution and presumed function of different cell types (Perez et al. 1999; Perez 2000).

7.4.5.1 Oral cavity and esophagus The first unit extends into the cephalic region where prey items are likely submitted to mechanical and enzymatic actions. It consists of the oral cavity (also interpreted as a pharynx connecting the mouth to the esophagus by Arnaud et  al. 1996; see also Figs.  7.3  A, B, 7.28  A, D, 7.29  A, and 7.36  A–B in this chapter) and the esophagus (Figs. 7.3 A, 7.19 A, 7.21 B, 7.28 A, D, 7.29 A, 7.31 A, 7.36 C–F, and 7.38). A prominent layer of cross-striated myofibrillous apparatus consisting of an inner layer of circular muscles sandwiched by longitudinal muscles surrounds the esophagus (Fig.  7.3  A, B). These muscles likely help to force prey into the intestine (Duvert & Salat 1995). The digestive epithelium of the cephalic region is composed of three columnar cell types (S1-, S2-, and S3-cells, compare

Fig.  7.36 in this chapter) exhibiting typical features of secretory cells, i.e., well-developed rough endoplasmic reticulum, Golgi bodies, and secretory granules gathered in the upper region of the cytoplasm. According to the ultrastructure of these secretory granules and the presence of coated vesicles which function to harbor glycoproteins, it has been hypothesized that secretory cells of the cephalic region are specialized either for the synthesis of mucosubstances (S1), or enzymes (S3), or both (S2) (Arnaud et al. 1996; Perez et al. 1999).

7.4.5.2 Intestine and rectum The second functional unit of the digestive tract extends into the trunk region and represents 90% of the gut length. It consists of a long and straight intestine (Figs.  7.2  A, 7.3  C, 7.29  A, 7.31  E, 7.34  C, and 7.42  A–B), which ends in a short vertical rectum ventrally connected to the anus just before the posterior septum (Fig.  7.2  A). Cilia are present both in intestinal and rectal cells (Fig. 7.37 C). The intestine and the rectum are surrounded by the visceral extracellular matrix (VECM) and a thin adjacent layer of coelothel, which consist of myoepithelial cells containing bundles of myofilaments whose orientation is mainly circular (Figs. 7.37 A, D, F and 7.39 C–F; see also Duvert & Salat 1995). As the equipment of myofilaments appears massive if compared with somatic myoepithelial cells (e.g., compare Fig. 7.40 G and Fig. 7.40 H), this layer may be considered a true circum­intestinal (= “peri-intestinal”) musculature. A complex system of flattened peritoneocytes may also surround the intestine and accompanied circum­ intestinal muscle cells. In S. cephaloptera, discontinuous peritoneocytes are often observed to fill small gaps in the line of circumintestinal muscle cells, where they wrap myelinated axons of the stomatogastric nervous system (Fig.  7.40  F–G). The circumintestinal muscles produce helicoidal peristaltism, inducing back and forth movement of the bolus into the intestinal lumen. Shinn (1997) also suggested that this anterograde and retrograde peristalsis could move the nutrient-rich hemal fluids located in the peri-intestinal sinus (Fig. 7.34 D) through the posterior sinus toward the ovarian space and the tail coelomic cavity. The anal sphincter consists of myoepithelial cells located at the epidermal level (Duvert & Salat 1995). Duvert and Gros (1982) mentioned the occurrence of five distinct junctional complexes between intestinal cells according to their apicobasal distribution: apical zonula adherens, paired septate junctions, including specific “pleated septate” junctions and “tricellular” junctions, and basal gap junctions. Hemidesmosome-like junctions are lacking.

7.4 Histological structure, cytology, and functional significance of organ systems 

The intestinal epithelium displays two categories of columnar ciliated cell, a fourth secretory type (S4-cell) containing large mucuslike granules, and an absorptive type (A-cell), which predominates in the posterior half of the intestine (Fig. 7.37 A). Several species have two intestinal diverticula in the most anterior part of the intestine, the epithelium of which is essentially composed of S4-cells (Arnaud et  al. 1999). The mucus-like secretion of the S4-cells likely participates in the lubrication of the bolus during peristaltic movements and/or in the formation of the peritrophic membranes surrounding the undigested nutrients (Reeve et al. 1975; Arnaud et al. 1996; Perez et al. 1999; Perez 2000). Such suggested double role is supported by the rapid necrosis of S4-cells observed in the absence of an alimentary bolus during starvation experiments (Perez et  al. 2000). The nutrients predigested in the lumen by the putative enzymes secreted by the cells of the cephalic region are endocytosized in the A-cells (Fig. 7.37 B). Indeed, although small electron-dense secretory granules are also gathered in the apical part of their cytoplasm, the A-cells exhibit organelles usually involved in the endocytosis of macromolecules associated with intracellular digestion, i.e., coated pits and endocytotic-coated vesicles, cytoplasmic tubules, and large vacuolar compartments containing heterogeneous material that probably correspond to endosome-like, endolysosome-like, and more terminal lysosome-like compartments involved in acid hydrolysis of macromole­cules (Fig. 7.37 B). The use of ferritin and peroxidase as tracers demonstrated that these organelles and vacuoles are involved in the endocytotic pathway (Perez 2000). The occurrence of intracellular digestion was evaluated in the intestine of P. elegans. Acid phosphatase histochemistry demonstrated the activity of this important and universal enzyme of the lysosomal compartment (Welsch & Storch 1983b). The basal cytoplasm of the A-cell is also occupied by large secretory granules and endocytoticcoated vesicles attesting for exchanges of macromolecules with the hemal system located in the subjacent visceral ECM and within the posterior septum (Fig.  7.37  D–E). Shinn (1993, 1997) suggested that the visceral ECM may be important as a route for metabolites from intestinal epithelium. Hemal fluids may circulate through the entire body driven by peristaltic contractions of the circumintestinal musculature. Exocytosis and endocytosis with coated vesicles at the basal pole of absorptive intestinal cells point to exchanges between the intestinal epithelium and the hemal system (Perez 2000). Ultrastructural changes of the intestinal epithelium have been described in fed and starved specimens of S.  cephaloptera (Perez et  al. 2000). Endocytotic vesicles arising from coated pits and endosome-like vacuoles

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containing electron-lucent material develop in the A-cells 5 min after the ingestion of a prey. During the following hours, large lipid inclusions, frequently observed in close contact with rough endoplasmic reticulum, accumulate in the apical cytoplasm and endolysosome-like compartments containing electron-dense heterogeneous material progressively arrange in columns according to the time of their formation, the youngest vacuoles localized at the apex, and the oldest at the basis of the cell. The development of light areas inside the dense contents of the older lysosomes could correspond to the utilization of digested material by the cell, or to its transfer to other tissues as ovaries for vitellogenesis. Because chaetognaths are devoid of specific storage structure, it is likely that the micromolecules obtained via intracellular digestion rapidly reach the hemal system located in the visceral ECM before to be delivered to the entire organism (Shinn 1997; Perez 2000). The intestinal epithelium undergoes marked ultrastructural changes during starvation experiments. In S4-cells, necrotic features appear and consist in a total disintegration of the organelles involved in the synthesis-secretion pathway. In A-cells, the endocytotic vesicles and endosomal-like compartment are larger than in fed specimens attesting to an increase in the membrane surface involved in endocytosis. The implication of receptors in absorption was evidenced by the presence on the vesicle membrane of a clathrin-like coat on the outer side and an electron-dense layer closely disposed against the inner side. Lipid-like inclusions and electron-dense vacuoles are absent. Although absorption is undoubtedly active beyond the 20th day of starvation, the intracellular digestive process is less evident because the endolysosome-like, electron-dense vacuoles are never observed (Perez et al. 2000). However, it is likely that intracellular digestion is functional in these A-cells because the fluid of electron-lucent vacuoles has a pH about 6.0 corresponding to the pH usually observed in endolysosome, and it contains high concentrations of NH4 and amino acids (Bone et al. 1987a), which certainly arise from intracellular digestion. Thus, the absence of dense vacuolar material does not reflect the absence of an intracellular digestion but rather a low concentration of absorptive products during starvation experiments (Perez et al. 2000). Observations of living specimens of S. cephaloptera showed that chaetognaths intermittently swallow seawater so that the continuous beating of cilia creates a water flow within the intestinal lumen even if it does not contain any food (Y. Perez, personal observation). Thus, during a short period of starvation, dissolved organic molecules certainly enter the intestinal lumen via this water uptake and could be internalized by the A-cells via the

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7.4 Histological structure, cytology, and functional significance of organ systems 

permanent receptor-mediated endocytosis. The irreversible necrosis of the intestinal epithelium within 1 month suggests that S. cephaloptera is poorly adapted to starvation during mating and reproduction (Perez et al. 2000).

7.4.5.3 Vacuolated intestine The cytological composition and anatomy described above seem highly homogeneous across the phylum, at least for the well-studied species belonging to the Sagittidae (including P. draco), Eukrohniidae, and Spadellidae families (Perez 2000). Nevertheless, the presence of an unusual, vacuolated intestine that obliterates the trunk coelomic cavity has been noticed in nine Sagittidae species (Dallot 1970). Bone et  al. (1987a) observed in P.  elegans that the large vacuoles contained high concentrations of free amino acids and NH4+. These authors showed this species to have neutral buoyancy with a lower specific density than P. setosa, a species without intestinal vacuo­ lation. It has been hypothesized that the replacement of heavy ions such as Na+ by NH4+ enhances the buoyancy of the vacuolated species by decreasing their specific gravity. Two types of vacuolated intestine have been recog­nized according to the number of vacuolated A-cells and the degree of vacuolation, e.g., the monoserial and the polyserial types (Dallot 1970) (Fig.  7.38). Ultrastructural data increased our understanding of the two types described previously (Perez et al. 2001). The development of the vacuoles is inversely proportional to the number of vacuolated A-cells, from the minimum of a single and large vacuolated A-cell on both lateral sides of the intestinal epithelium as in M. minima and D. decipiens to up 10 to 20 smaller vacuolated A-cells as in P. megalophthalma and S. zetesios, respectively (Fig. 7.38). The intestinal vacuolated cells represent cells of the same type as absorptive A-cells. Vacuolated intestinal cells always exhibit the organelles implicated in endocytosis of macromolecules, and the large vacuoles result from an increment in volume of the endolysosomal compartment (Perez et al. 2001). The implication of the large vacuoles in the intracellular digestion process is supported by the acid pH of the vacuolar fluid and the storage of NH4+ and free amino

 245

acids. NH4+ ions could arise from intracellular digestion of macromole­cules, and its concentration might be quickly adapted to allow the vacuolated species to move and hunt without effort in a wide range of depth. The development of large vacuoles appears as a cytological adaptation to deep and cold water environments that consists in a tradeoff between the energy allocated to locomotion, feeding, and reproduction (Perez et al. 2001).

7.4.6 Body cavities Chaetognaths show several body cavities which, from typological point of view, can be distinguished into two classes: (1) primary body cavities lined by an ECM secreted by surrounding epithelial cells with their apices turned away from the cavities and (2) secondary body cavities, called true coeloms, lined by a mesoderm-derived epithelium (called coelothel) with the cells’ apices facing the cavities (as defined by Schmidt-Rhaesa 2007).

7.4.6.1 Primary body cavity—hemal system Primary body cavities may be remnants of the persisting blastocoel or derive from secondary expansion of intercellular spaces in various mesodermal tissues (Ruppert 1991; Schmidt-Rhaesa 2007). In chaetognaths, primary body cavities are only visible in early embryogenic stages and vanish completely shortly after gastrulation takes place (Kapp 2000). In adults, newly derived equi­ valents to primary body cavities do occur but are small and hard to identify, as they are restricted to slightly widened compartments of the visceral extracellular matrix (VECM). In F. hispida (Sagittidae), the ECM of the circumintestinal muscle (myoepithelial) cells looks widened; the basal laminae of the intestinal, S-type epithelial cells and circumintestinal muscle cells enclose an amorphous, extremely electron-dense lumen, the peri-intestinal sinus, that is considered part of a hemal system (Shinn 1993, 1997: figs. 59, 60B, 61; and Fig. 7.34 D in this chapter). Peri-intestinal lumina are most voluminous mediodorsally and medioventrally, namely, at

◂ Fig. 7.36: Chaetognath digestive system in the cephalic region. Light (A) and TEM (B–F) micrographs in Serratosagitta serratodentata (A, B, D–F) and Parasagitta setosa (C). A, Section through the oral cavity. Arrowhead indicates the cuticularized epithelium which occurs ventrally on the head. B, Detail of the apical cytoplasm of S1-(S1), S2-(S2), and S3 (S3)-cells identifiable by the fine structure of their secretory granules. C, Section through the esophagus. In the esophageal S2-cells, the secretory granules undergo a maturation of their central core into a network of less dense and heterogeneous material. D, Detail of an immature secretory granule of S1-cells close to the trans-Golgi network. E, Detail of two mature secretory granules of S1-cells gathered in the upper part of the cytoplasm. The dense core exhibits a paracrystalline structure. F, Detail of mature secretory granules of S3-cells. Note the denser central zone and the irregular outline of the granules (modified from Perez et al. 1999). Further labels: es, esophagus; mo, mouth opening; ph, pharynx.

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7.4 Histological structure, cytology, and functional significance of organ systems 

 247

Fig. 7.38: Sections showing a monoserial (A) and polyserial (B) type of vacuolated intestines. Original light micrographs (semithin sectioning histology). A, Decipisagitta decipiens; B, Parasagitta megalophthalma. Labels: Asterisk, vacuolated intestinal cell; int, intestine; ov, ovary; lf, lateral fin; tlmu, primary longitudinal (trunk) musculature; trco, trunk coelom.

the contact zone of the VECM and dorsal and ventral mesenteries (compare fig. 60A in Shinn 1997). Posteriorly, the peri-intestinal sinus passes into paired saccate spaces, called the posterior sinuses (Shinn 1997), which are bordered by the intestine (medially), the tail coelom (posteriorly and dorsally), the trunk coelom (laterally and ventrally), and the peri-ovarian coelom/space (laterally). The posterior sinuses are traversed by a complex meshwork of ramifying, non-epithelial cells, some cytoplasmic projections adjoin the posterior septum. The fenestrated, podocyte-like appearance of the myoepithelial cells establishing the posterior septum tail side let Shinn (1993, 1997) conclude that hemal fluids, potentially carrying dissolved metabolites from the intestine and pushed posteriorly by peristaltic movements of circumintestinal muscles, may be extruded into the tail coelom by ultrafiltration. However, preliminary observations on comparable intestinal surroundings in S. cephaloptera do not reveal a hemal system at all. The VECM is thin and does not show a peri-intestinal sinus structure between its basal laminae (compare Fig.  7.40  C–G). Moreover, the structural equivalent to paired posterior sinuses more resembles features of additional, postovarian coelomic cavities (see section below). The ECM of these sinuses is not continuous with the VECM. Therefore, it has to be doubted

whether a hemal system is common in Chaetognatha. Nevertheless, the existence of a putative open circulatory system from the remainders of the primary body cavity (Shinn 1993, 1997) has important implications because the anatomical position of these sinuses resembles the organization of the annelid circulatory system (Perez 2000; Malakhov & Berezinskaya 2001). Malakhov and Berezinskaya (2001) noticed that chaetognaths lack the ventral vessel that most annelids possess and concluded that their circulatory system is simpler with respect to the complete absence of endothelial-like tissue and its general organization.

7.4.6.2 Secondary body cavities—trimeric coelomic organisation The secondary (coelomic) body cavities in chaetognaths show a trimeric organization in adults. Three body parts are established comprising enigmatic paired head coeloms as well as obviously paired trunk and tail coeloms. By contrast, only two body parts can be distinguished during early embryogenesis. This bimeric organization includes paired head coeloms and precursors of paired trunk coeloms (Kapp 2000). All coelomic cavities are lined by columnar, cuboidal, or flattened coelothelial

◂ Fig. 7.37: Chaetognath digestive system in the trunk region. TEM micrographs of the intestinal epithelium in Sagitta bipunctata (A, E), Serratosagitta pacifica (B), Parasagitta setosa (C), and Serratosagitta serratodentata (D, F). Originals. A, General aspect of the intestinal epithelium and the surrounding peritoneum showing the secretory (S-4) cells with well-developed endoplasmic reticulum (rER) and absorptive (A) intestinal cells. B, Observe the apical vacuole (v) of an A-cell surrounded with a well-developed tubulovesicular network, the subjacent large vacuoles with dense bodies (db), and the relative scarcity of apical secretory granules (arrow) gathered just below the apical membrane. C, Detail of the apical membrane of an A-cell. Arrow heads indicate two coated vesicles in formation (coated pits) attesting for a receptor-mediated endocytosis of macromolecules. Note also the cilia (ci) in the intestine lumen. D–E, Detail of the basal cytoplasm of an A-cell with a secretory granule (asterisk) (D) and a clathrin-like coated endocytic vesicle formation (arrowhead) (E) indicating exchanges between the A-cells and the hemal system situated within the visceral extracellular matrix (vecm) surrounding the intestinal epithelium. F, Myoepithelial cells of the coelothel located around the intestine. Observe the myofilaments (myf) and large Golgi body (go). Further labels: intl, intestine lumen; mi, mitochondrion; myc (gmu), circumintestinal myoepithelial cells/gut musculature; trco, trunk coelomic cavity.

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cells which, with only one exception, contain myofilaments. The somatopleura mainly consists of crossstriated, locomotory muscles with transverse, oblique, or longitudinal orientation (e.g., Figs.  7.3, 7.39  A–B, 7.41  A, 7.42  A, and 7.45  A–B; for details, see descriptions in Section 7.4.4, “Muscular Apparatus and Locomotion”). Along the mediosagittal and mediolateral plane, namely, between the head muscles as well as in the contact zones of two adjacent pads of the primary longitudinal muscle in trunk and tail, somatic myoepithelial cells are observed (ciliary myoepithelial lateral field cells, medial myoepithelial cells, perimysial cells). These myoepithelial cells lack sarcomeres but contain numerous, more or less aggregated myofilaments located in basal position and running parallel to the ECM (e.g., Fig. 7.40 F, H). The lateral field and medial myoepithelial cells sitting on the epidermal ECM may taper into small apical processes wrapped upon each other. In addition, their lateral cell borders intertwine and are firmly connected by septate junctions (Fig. 7.40 H). This results in firm adhesion and strength and enables the myoepithelium to work against the coelomic pressure. Squamous peritoneocytes are another type of mesodermal epithelial cells that may occur in all coelomic cavities. They strongly vary in shape and thickness but mostly display flattened profiles (compare Figs. 7.39 C–E, 7.40  F, and 7.41  B–E). The cytoplasm contains numerous membrane-coated tubules and granules with presumably secretory contents of high electron-density, is riddled with aggregated microtubules, and is always devoid of myofilaments (Figs. 7.39 C–E, 7.40 F, and 7.41 E). The formation of peritoneocytes is epithelium-like proximal to the locomotory (including the primary longitudinal) musculature (Fig. 7.39 C) but remains fenestrated in the area of the lateral field cells and along medial (mesenterial) border of the trunk and tail coeloms (Figs. 7.39 D–E and 7.40 F, see further description and typological discussion below). In S. cephaloptera, loosely aligned peritoneocytes exclusively provide a fenestrated epithelium, which forms an ECM with attached myepithelial cells of the trunk, peri-ovarian, and tail coeloms as well as with the circumintestinal myoepithelium (Fig. 7.41 B–E). Scattered peritoneocytes may fill in gaps left by the circumintestinal myoeithelial cells. Occasionally, flattened ramifications of these peri-intestinal peritoneocytes cover myelinated axons of stomatogastric motor neurons (Fig. 7.40 F). The origin of the peritoneocytes remains unclear. Neither in gastrulating embryo nor in the hatchling these cells can be attributed to components of the developing coelomic epithelium (Shinn & Roberts 1994; Shinn 1997). The formerly paired head coelomic cavities are somehow scattered in the adult chaetognath body as

medial mesenteries are missing. Instead, head coeloms are restricted to several interspaces of various head muscles, the (peri)esophageal musculature, and the anterior septum (Figs. 7.19 A, 7.20 A, and 7.39 A–B). Both trunk and tail include two large, bilaterially symmetrically arranged pairs of coelomic sacs, each pair separated from each other by medial mesenteries (Fig. 7.39 A, D–E). Both coelomic cavities in the trunk occupy the interspace built by the (1) intestine (medially); (2) oblique, transverse, and (primary) longitudinal locomotory muscles (dorsallylaterally-ventrally); (3) the anterior septum (anteriorly); (4) the posterior septum (posteriorly); and (5) the ovaries (bulging deeply into the trunk coeloms). Ventro- and dorsomedially of the intestine (along mediosagittal plane), the trunk coelomic compartments are separated by the ventral and dorsal trunk mesenteries, consisting of the mesenterial ECM and two adjacent layers of myoepithelial (coelothelial) cells. The mesenterial ECM tapers and subsequently disappears when approaching the epidermal ECM at the level perimysial epithelial cells (cf. Fig.  7.35  A, E). The dorsal trunk mesentery also separates both ovaries (Fig. 7.39 A, D–E). Each ovary is embedded into an independent coelomic cavity, termed here as peri-ovarian (coelom) space (Fig.  7.40  A–E). The various maturation stages of aggregated oogonia and oocytes (see Figs.  7.1  B, C, 7.40, and 7.45  A, C) almost completely occupy the ovaries and therefore leave little space for the peri-ovarian coelom, which is bordered to its outer face by the ovarian wall and to its inner face by the follicular reticulum encompassing the oogonia and oocytes (e.g., Fig.  7.40  B, D). There is no direct connection between the female gonopore, the associated oviduct, and the peri-ovarian coelom. Sperm transfer and release of fertilized eggs have to happen through the medial wall of the oviducal complex (for details, see Section 7.4.7.2, “Ovary and Oviducal Complex” and Section 7.5.1, “Mating, Fertilization, and Egg Laying”). Posteriolateral of the intestine and posterior of the ovaries, a pair of small cavities, filled by an extremely electron-dense, amorphous matrix which looks similar to the surrounding trunk, peri-ovarian and tail coelomic fluids, is lined by fenestrated epithelium of peritoneocytes. Based on their relative dense packing and orientation relative to the subjacent ECM (apex facing the cavity’s lumen), the fenestrated epithelium of peritoneocytes may be considered coelothelial cells and, subsequently, the enclosed space a true coelom, however small. These postovarian coelomic spaces in S. cephaloptera are most likely not homologous to the posterior sinus described in sagittid chaetognaths (Shinn 1993, 1997) because here the postovarian spaces do not seem to have any spatial connection to the VECM (Fig. 7.41 B).

7.4 Histological structure, cytology, and functional significance of organ systems 

Both coelomic cavities in the tail are the most voluminous in the chaetognath body. Medially, both cavities are separated by the medial mesentery composed of a mesenterial ECM and cuboidal (medial) myoepithelial cells at either side (Figs. 7.39 A and 7.42 A–B). In sagittid chaeto­ gnaths, those myoepithelial cells lining the anterior aspect of each tail coelomic cavity may strongly ramify into many slender, interdigitating cytoplasmic extensions. Shinn (1997) emphasized structural and probably also functional correspondences of these fenestrated tail myoepithelial cells to podocytes present in some bilaterian extretory organs (compare his figs. 61 and 62). However, S. cephaloptera shows myoepithelial cells at anterior end of the tail coeloms to be columnar and heavily folded at their bases (Fig. 7.40 F). This subcellular pattern is reminiscent of a basal labyrinth typically found at base of secretory or transporting epithelial cells. The tail coelom contains numerous, aggregates of successional spermatogenic stages deriving from paired testes (column of densely packed spermatogonia) nested in specialized lateral field myoepithelial cells (Bergey et  al. 1994; Shinn 1887). The initially syncytial spermatogenic masses float through the coelomic space/fluids (for details, see Section 7.4.7.1, “Testis and Seminal Vesicle”). Mature sperm or spermatophores are discharged via paired coelomoducts each of which is opens into the tail coelom lateroposteriorly. Original TEM data first presented in this book chapter now clearly reveal that the distal portions of coelomoducts (spermioducts) open into the tail coelom via a ciliated funnel (see Fig. 7.43), made up by multiciliated peritoneal epithelial cells. These coelomoducts are closely similar to nephrostomes of metanephridial organs as well as to transformed gonocoelomoducts, respectively. All coelomic fluid contains high levels of free amino acids. Its composition is very similar to seawater with respect to Na+, Cl−, SO42−, NH4+, and Mg2+ concentrations (Bone et al. 1987a). In Parasagitta setosa, 60% of the free amino acids are glycine and alanine. Ultrastructural observations of the mesoderm subdivision and coelomogenesis have been well documented in hatchlings of the planktonic species F. hispida (Shinn & Roberts 1994; Shinn 1994a, 1997) and the benthic species S. cephaloptera (Harzsch et al. 2015).

7.4.6.3 Status and phylogenetic significance of chaetognath coeloms and associated metanephridial organs The general body cavities of the trunk and tail are considered to be a true coeloms lined by a continuous coelothel

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(Duvert & Salat 1979; Welsch & Storch 1982; Shinn & Roberts 1994; Shinn 1994a, 1997; Harzsch et  al. 2015). However, coelomogenesis, called heterocoely, is unique in chaetognaths and may therefore be considered apomorphic for this taxon (Kapp 2000; Perez et  al. 2014). Head coeloms are developed differently from trunk and tail coeloms, alike anterior and posterior septa the latter of which derives from interstitial coelothelial cells crammed between primordial germ cells (for details, see Section 7.5.2, “Germline, Cleavage, Gastrulation, and Coelomogenesis”). Therefore, it remains ambiguous which chaeto­ gnath coeloms or at least parts of their coelomic systems may be compared and potentially homologized to equivalent coeloms in other invertebrate taxa with bimeric or trimeric body organization. After all, ultrastructural, topological, and functional correspondences are indicated in the gonocoelomic cavities and gonocoelomoducts of many lophotrochozoans and chaetognaths. The coelomoducts at posteriolateral ends of the chaetognath tail may mainly act as spermio­ ducts, indicated by the presence of sperm-storing seminal vesicles at the proximal end of the spermioducts and thereby, for example, resemble the gonocoelomic system of mollusks. Whether tail coelomoducts of chaetognaths also divert excretion is a matter of discussion. At least, Shinn (1997) documented in the sagittid species F. hispida a fenestrated myoepithelium lining the tail coelom anteriorly adjacent to the posterior sinus (Fig.  7.61, p. 175). If these epithelial cells were podocytes, ultrafiltered excretory products would be able to enter the coelomic cavities of the tail and leave the body through the presumed gonocoelomoducts as well. Spatial and functional combination of gono- and renocoelomic systems would somehow integrate characteristics known from mollusks (gonocoelom) and annelids (metanephridial organs). However, annelid metanephridia are transsegmental and develop differently in comparison with isosegmental tail coelomoducts of chaetognaths. Reports on putative coelomoducts associated with body parts others than the tail do exist for chaetognaths but remain preliminary until fine-scale TEM studies will have been conducted. In few basally branching genera, i.e., Archeterokrohnia, Heterokrohnia, Eukrohnia, and Bathyspadella, there are reports on a pair of epidermal glandular canals which are assumed to open posteriorly on the dorsolateral side of the neck as well as anteriorly into the scattered compartments of the head coeloms (Tokioka 1939; Hyman 1959; Kuroda 1981; Casanova 1991a, 1992; Casanova & Chidgey 1987; Shinn 1997). These ducts may represent coelomoducts, as well. However, data on their ultrastructure and their involvement in a putative excretory function are not available

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Fig. 7.39: Diversity and location coelomic cavities in chaetognaths exemplified by means of original histological (A–B, stained with toluidine blue) and TEM micrographs (C–E). Spadella cephaloptera. A, Mediohorizontal (longitudinal) semithin section showing minute coelomic cavities within the head (hco) posterior to the external vestibular dilator muscle (musculus dilator vestibuli externus = mdve) as well as more spacious coelomic cavities in the trunk (tco) at either side of the intestine (int), with paired peri-ovarian coeloms (pco) included, and most voluminous coelomic cavities in the tail (taco) containing the spermatogenic masses of the testes (te). Note the dorsal mesenterial septum in the trunk (dtme) as well as the mesentery (tamu) dividing both testes coeloms in the tail. B, Mediosagittal semithin section complementary to A showing various compartments of the tail coelom located ventral (vtco) and dorsal (dtco) of the intestine and ovaries. C, Horizontal section of peri-esophageal head coelom cavity close to transition zone to the trunk; section area is indicated by dashed box

7.4 Histological structure, cytology, and functional significance of organ systems 

so far. If this hypothesis is confirmed by new data, it will have important impact on our understanding of the phylogenetic relationships of chaetognaths. More specifically, it will be important to analyze if these ducts are parts of a proto- or metanephridial system because the homology of protonephridia among lophotrochozoan taxa is now widely accepted (Bartolomaeus & Ax 1992; SchmidtRhaesa 2007; Baeumler et al. 2012). These future analyses including biochemistry and cytophysiology may reveal if the tail coelomic cavities and associated coelomoducts of chaetognaths really perform two functions (extretion, accommodation of spermatogenic masses, and discharge of sperm) simultaneously. If such dual function is confirmed, reno-gonocoeloms with nephrostome-equipped coelomoducts should be evaluated as a potential, longsought character to ally chaetognaths with other bilaterians. Similar coelomic systems coupling excretion and reproduction are found in metacoel of various lophotrochozoan ingroups, such as sipunculans (Rice 1993), echiurids (Pilger 1993), phoronids (Herrmann 1997), and brachiopods (James 1997; Lüter 2016). Admittedly, coelomic cavities and associated nephridial organs are considered structures of low complexity what generally hinders attempts to convincingly support primary homology hypotheses (Koch et al. 2014). Koch et al. (2014) referred to coeloms and nephridia of annelids and panarthropods and stated that there are only three reliable, independent characters to homologize these organs in both taxa. However, the specific spatial coherence and functional coupling of gonads, coelomic cavity, podocyte-based excretion, and bifunctional coelomoducts may at least sufficiently increase aspects of structural complexity, which makes it worthwhile to reassess the chaetognath tail coelom in a phylogenetic context. In case that the Hyponeuria concept was valid (see Perez et al. 2014; and Fig. 7.6 in this chapter), one option would be to consider chaetognaths to be the sister group or a basal ingroup of Lophotrochozoa. Then reno-gonocoeloms with nephrostome-equipped coelomoducts may be interpreted a synapomorphy of this grouping. However, because of unresolved rooting of Protostomia, the Hyponeuria concept currently also allows for placing Chaetognatha as sister group of Protostomia (e.g., Marletaz et al. 2006 and Fig. 7.6

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in this chapter). Then, the assumption of convergent evolution of reno-gonocoeloms with nephrostome-equipped coelomoducts becomes more likely. The latter assumption would fit the generally accepted view that in light of the Ecdysozoa/Lophotrochozoa hypothesis, for example, metanephridial organs have evolved at least three times independently among Bilateria (see summary in SchmidtRhaesa 2007).

7.4.7 Reproductive system The male (Figs. 7.42–7.44) and female (Fig. 7.45) reproductive organs may occupy more than half of the body volume and lie dorsally at the anterior and posterior sides of the posterior septum (see Fig.  7.1). An unusual genital structure described in Heterokrohnia (Casanova 1985a) consists of an “annex secretory gland” situated around the intestine at the level of the posterior septum. This structure opens within the ovaries and connects the ovarian spaces and the tail coelomic cavity, a situation that recalls the hermaphrodite gland of mollusks (Casanova 1985a). However, neither histological nor cytological data are available to unequivocally support such an anatomical connection between the female and the male reproductive system. Shinn (1997) suggested that owing to its anatomical position and continuity with the posterior septum, this gland could be part of the hemal system. The morphology and histology of the reproductive organs in chaetognaths have been described in S. bipunctata (Hertwig 1880; Grassi 1883; Stevens 1903, 1905, 1910; Burfield 1927; Ghirardelli 1968), P. elegans (Stevens 1905, 1910), S. cephaloptera (Ghirardelli 1954a, 1956, 1961a, 1968), P. draco (Ghirardelli 1953), F. enflata (Ghirardelli 1954b), P. lyra and F. hexaptera (Ghirardelli 1961b, 1968), S. planctonis (Pierrot-Bults 1975b), and S. serratodentata (1976b). Several aspects of the reproduction and reproductive organs have been reviewed by Ghirardelli (1968), Reeve and Walter (1972b), Alvariño (1983, 1992), Reeve and Cosper (1975), Strathmann and Shinn (1987), Kapp (1991a), Pearre (1991), and Shinn (1997). The following anatomical and cytological descriptions are mainly based on the extensive studies of Shinn (1992, 1994b, 1997) and Bergey et al. (1994) who thoroughly

◂ in A. Lateral myoepithelial cells (lateral field cells = lfc) line the somatic aspect of the cavity; visceral aspect is represented by medial myoepithelial cells (indistinct on image) and intertwining overlayer of granulated peritoneocytes (per). D, Close-up of (dorsal) mesenterial septum of the posterior part of the trunk (trme) crammed between both ovaries. E, High-power magnified dorsal, interovarian mesentery comprising a median extracellular matrix (mecm) lined at either side by mesenterial myoepithelial cells (mec) and locally by incompletely overlaying granulated peritoneocytes. Coelomic spaces are restricted to narrowed interlayers between mesentery and the myoepithelial cells making up the ovarian wall (ow); myofilaments of ovarian wall cells are marked by arrowheads. Further labels: a, anus/rectum; cc, corona ciliata; cfo, ciliary fence organ; cu, cuticle; eg, fertilized egg; es, esophagus; fgp, female gonopore; gs, grasping spine; ho, hood (praeputium); mep, multilayered head epidermis; ocp, previtellogenic oocyte; odc, oviducal canal cells; oo, oolemma; ov, ovary; ovc, oviducal complex; pec, proximal epidermal cells; ps, posterior septum; tlmu, primary longitudinal (trunk, tail) musculature; sg, secretory granules; vec, vacuolated epidermal cells of the collarette region.

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studied the cytoarchitecture and functional organization of the female and male reproductive systems in F. hispida.

7.4.7.1 Testis and seminal vesicle Each testis consists of syncytial spermatogonia accommodated in both coelomic cavities of the tail (e.g., Figs. 7.35 A and 7.39  A). Tail coelom and spermatogonial masses at various stages of succession are lined laterally by a flattened somatic sheath made of specialized unciliated lateral field cells and partially covered by thin extensions of peritoneocytes, as well as medially by medial mesodermal epithelial cells (Bergey et  al. 1994; Shinn 1997; see also Fig.  7.42  B in this chapter). Spermatogonia start their development in the testes and are released into the tail coelomic cavities from the anterior ends of the testes (Figs. 7.32 E, 7.35 A, and 7.42 A–D). They remain syncytial and, by mitosis and meiosis, give rise to spermatocytes and elongated spermatids, and finally to differentiate into spermatozoa, which gradually fill the tail coelomic cavities such that each cavity contains spermatozoa at different stages of development in mature specimens (Figs.  7.32  E and 7.42  E). Developing spermatogonia and spermatozoa are maintained in a continuous circular movement by the activity of ciliated cells associated with two median mesenteries, which divide partially into two adjacent compartments each becoming a tail coelomic cavity (John 1933; Alvariño 1983; Bergey et al. 1994; Shinn 1997). According to Deurs (1972, 1875), each filiform spermatozoon of S. cephaloptera carries a centriole-like structure close to the anterior end from which a typical axoneme projects. Further conspicuous organelles found

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in spermatozoa are a single and stretched mitochondrion, a banded vesicle consisting of alternating vertical layers of different electron density, and a likewise elongated nucleus with extremely condensed caryoplasm (see also Fig. 7.42 F, G in this chapter). Other, membrane-lined organelles, such as a membranous cap acting as acrosome or beaded structures, may occur, as well. Ultrastructure of the filiform spermatozoa of Parasagitta euneritica and F. hispida turned out to be very similar to those of S. cephaloptera (Alvariño 1983; Shinn 1997). Two spermioducts (syn.: sperm ducts, vasa deferentia) are found posteriorly, one on each side of the tail, and connect the coelomic cavities with the seminal vesicles. The anterior tip of the sperm ducts looks like a ciliated funned resembling a nephrostome associated with metanephridial organs (see Fig. 7.43 A and discussion in Section 7.4, “Histological structure, cytology, and functional significance of organ systems”). The outer face of the ciliated funnel is elaborated from lateral peritoneocytes, its inner face as well as anterior portion of the sperm duct are lined by cuboidal, multiciliated epithelial cells the cytoplasm of which contains numerous apical microfilaments and mitochondria, Golgi bodies, and vesicles with an electrondense core. At the tip of the apex, at least three, vibratile, and elongated cilia are protruded (Fig.  7.43  B), which invade the sperm duct and found in close aggregations (Fig. 7.43 C). Distal and median sperm duct cell differ from multiciliated funnel epithelial cells by a massively developed cytoskeleton of bundled tonofilaments attached basally to peg-like protuberances of the epidermal ECM (Fig. 7.43 D–E). Ciliary equipment is not obvious in this cell type. The median duct lumen is often observed to be compressed (Fig. 7.43 D). The proximal sperm duct epithelium

◂ Fig. 7.40: Histology (A, stained with toluidine blue) and TEM-based ultrastructure (B–H) of peri-intestinal and peri-ovarian coelomic cavities in posterior half of the trunk of Spadella cephaloptera. A, Parasagittal LM semithin section showing the posterior end of the intestine the lumen of which bends ventrally and passes into the anal region (a). Compartments of both trunk coelomic cavities are seen above and below the intestine and ovaries (dtco, vtco). Note the occurrence of the posterior septum (ps) separating trunk from tail coeloms. B, Parasagittal section of dorsal trunk coelom (dtco) located below the (dorsal) multilayered epidermis (mep) and dorsal to the ovarian space (section region indicated by dashed box in A). It is lined dorsally by a unilayer of medial myoepithelial cells (myc) and ventrally by a diffuse double layer of polymorphous peritoneocytes (per) and epithelial cells of the ovarian wall (ow). The peri-ovarian coelom (pco) here forms a narrow interface between the ovarian wall and the only locally compact cell profiles of the follicular reticulum (fr). Parasagittal section. C, Parasagittal section of sheath-like, dorsal compartment of the trunk coelom crammed between the intestine (int) and an ovary, the follicular reticulum is minute, the ovarian wall epithelium is fenestrated in some places (arrowheads). Sector indicated by dashed box in A. D, Ovary-intestine interface in greater detail slightly posterior to C lacking the coelomic cavity but instead displaying closely adjoined profiles of myoepithelial cells containing numerous myofilaments (myf), the peri-ovarian coelom is compressed here to a tiny interlayer (arrow). Parasagittal section. E, Parasagittal section of posterior ovary region with abutting, moderately voluminous spaces of peri-ovarian and trunk coelom (sector indicated by dashed box in A), note the breakthrough in the ovarian wall (arrow) indicating partial porousness of both coelomic spaces. F–H, Sequence of parasagittal section showing the ventral aspect of the trunk coelom (vtco) with associated mesodermal epithelia: F, overview, note the locally inflated profile of a peritoneocyte surrounded a myelinated neuron (myn); G, close-up of dorsal lining of ventral aspect of trunk coelom achieved by visceral myoepithelial cells (myc (gmc)) establishing circumintestinal/gut musculature (gmu), sector marked by dashed box in F; H, columnar myoepithelial cells subjacent to epidermal extracellular matrix (ecm), lateral cell borders are strongly intertwined, sector indicated by dashed box in F. Originals. Further labels: cof, collagenous microfibrils; eg, fertilized egg; jec, jelly coat of vittelogenic oocyte; intl, intensinal lumen; mi, mitochondrion; ocv, vitellogenic oocyte; ooc, germinal cluster of various oogonia; owc, ovarian wall cell; pec, proximal epidermal cells; ps, posterior septum; tlmu, primary longitudinal (tail) musculature; te, testes; vecm, visceral extracellular matrix.

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7.4 Histological structure, cytology, and functional significance of organ systems 

houses both secretory cells containg electron-dense granules and especially to both lateral borders multiciliated epithelial cells (Fig.  7.44  C–D). The seminal vesicles, located on each side of the posterior lateral end of the tail, mostly consist of a simple glandular epithelium surrounding a more or less inflated lumen that is potentially completely occupied by mature spermatozoa (Figs. 7.43 F and 7.44  A–B). The seminal vesicle is embedded in the pluri­stratified (multilayered) epidermis whose secretions wrap the spermatozoa in a surrounding coat (Fig. 7.44 A, see also illustrations provided by Bergey et al. 1994; Shinn 1997). During mating, mature seminal vesicles are drained through a temporary slit along a preformed line bordered by specialized suture cells, and one or two sperm masses, naked or enclosed within spermatophores, are deposited on the mate. The mechanism of suture opening remains unknown at the cellular level (Shinn 1997).

7.4.7.2 Ovary and oviducal complex The paired ovaries are found dorsally in the posterior part of the trunk (Fig.  7.45  A–B) and separated from the trunk coelomic cavity by a thin retrocoelothelial somatic (ovarian) wall that consists of squamous myoepithelial cells the contractile apparatus of which is mostly circular, but laterally also longitudinal myofilaments can be observed (Shinn 1992, 1994b, 1997). Egg laying is presumably facilitated by the myoepithelial cell contraction. The myoepithelial cells enclose a fluid-containing ovarian coelomic subcompartment (termed here as peri-ovarian coelom) and differentiating oocytes (including previtellogenic, vitellogenic, and postvitellogenic oocytes) arising

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from female germ cells packed in the median side of the oviducal complex. The ovarian wall is mostly compact, although the myoepithelium looks interrupted in some places. There, the trunk and peri-ovarian coelomic cavities/subcompartments seem to be connected (Fig. 7.40 E). The larger size class of oocytes corresponds to fully grown oocytes with a large central nucleus typical of the aspect prior to ovulation (Fig.  7.45  C–D). A follicular reticulum encloses the oocytes and the dorsal and ventral secretory wings of the oviducal complex (Fig.  7.45  D). Follicular reticulum cells may appear extremely flattened and barely recognizable, especially in those sections taken from the ventral border of the ovaries. Accordingly, the peri-ovarian coelomic space may be diminished to a thin sheet along the inner face of the ovarian wall myoepithelial cells (Fig.  7.40  B–D). Its cytoplasm exhibits few synthetic organelles and is dominated by numerous vesicles. The plasma membrane of the follicular reticulum and those of the oviducal cells are connected by gap junctions. The cellular or syncytial organization of the follicular reticulum remains unknown (Shinn 1997). Shinn (1992, 1997) suggested a possible function of the follicular reticulum, including secretion of egg coat, transport of yolk precursors to oocytes. The ovaries do not directly open outside via a continuous oviduct and terminal female gonopore. Instead, an oviducal complex is present that establishes an epithelial barrier (“cellular sheath of the oviducal complex”: Shinn 1997) between the actual oviduct and the posterolateral periphery of the peri-ovarian coelom (see schematic reconstruction on fig. 77 in Shinn 1997). The oviducal complex consists of a anterolaterally closed duct and posteriorly opened at the level of the female genital openings and

◂Fig. 7.41: Histology (A, stained with toluidine blue) and TEM-based ultrastructure (B–F) of transition zone of trunk and tail documenting the posterior septum and small coelomic cavities/compartments of the postposterior trunk region of Spadella cephaloptera. A, Parasagittal LM semithin section. Details are described in caption of previous Fig. 7.40 A, sector indicated by dashed box in A. B, Enigmatic postovarian cavity, presumably coelomic (poco), located between dorsoposterior face of the intestine (int), the dorsal compartment of the trunk coelom (dtco), and the posterior septum (ps) and posteriorly adjoined coelomic cavities of the tail (taco). This presumed postovarian coelom can be identified by the presence of an extracellular matrix (ecm) and adjoined fenestrated epithelium lining its inner face. Flattened epithelial cells ramify into the coelom and are continuous with and hence termed as peritoneocytes (per) resting on myoepithelial cells (myc) sectorally lining the other coelomic cavities. C, Epithelial lining of presumed postovarian coelomic space in greater detail; note the outer overlay of myoepithelial cells containing numerous myofilaments (mif) and inner fenestrated layer of peritoneocytes, the associated extracellular matrix is clearly discernible by higher electron-density, sector indicated by dashed box in B. D, Close-up of part of the posterior septum abutting the presumed postovarian coelom, extracellular matrix adjoins the coelomic cavity directly over longer distance, most anterior myoepithelial cells of the tail display slender, microvilliform projections (arrow) but do not convincingly match podocyte characteristics, sector marked by dashed box in B. E, Ventral rim of presumed postovarian coelom in greater detail, two peritoneocytes meet but leave a gap (double arrows) where the coelomic fluids come in direct with the extracellular matrix, the peritoneocytes are re-inforced by microtubules (mic) and probably also by myofilaments, sector indicated by dashed box in B. F, Parasagittal section of more ventral aspect of the posterior septum showing complex packing of strongly intertwined myoepithelial cells lining small ventral branches of the trunk coelom (vtco), the extracellular matrix of the posterior septum contains pads of presumably collagenous microfibrils (cfp), associated most anterior myoepithelial cells lining tail coelom display multiply invaginated base resembling a basal labyrinth, podocyte characteristics are not obvious, sector indicated by dashed box in A. Originals. Further labels: a, rec anal/rectal region; eg, fertilized egg; gmu, circumintestinal/gut musculature; intl, intestinal lumen; mep, multilayered epidermis; nu, nucleus; ocp, previtellogenic oocyte; rER, rough endoplasmic reticulum; tlmu, primary longitudinal (tail) musculature; te, testes; vecm, visceral extracellular matrix.

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seminal receptacles. It is composed of an inner syncytium surrounded by an outer cellular sheath (Fig. 7.45 E–F). The oviducal syncytium appears to have no permanent lumen and contains elongate, anastomosing membrane-bounded saccules. Foreign spermatozoa stored in the seminal receptacles after mating extend into the space enclosed by these two concentric layers along the entire length of the oviducal complex (Fig.  7.45  F). Dorsal and ventral, flattened extensions of the oviducal cellular sheath consist of secretory cells with extensive rough endoplasmic reticulum, numerous Golgi bodies, and basal exocytosis of vesicles into the ovarian space. Central oviducal cells are circularly arranged and are connected to the surrounded oviducal syncytium cells by interdigitating microvilli and gap junction. Each previtellogenic oocyte is associated with two oviducal specialized cells, the so-called accessory fertilization cells (AFCs, see fertilization process).

7.5 Reproduction and ontogeny The embryonic development of chaetognaths is known from the key studies conducted by Hertwig (1880), Elpatievsky (1909), Doncaster (1902), Burfield (1927), and John (1933). Laboratory culture has been an essential tool for our understanding of their life cycle (reviewed in Harzsch et  al. 2015) and allowed studying various topics, including the effect of laboratory culture conditions and food on growth (Murakami 1966; Reeve 1970a, 1970b; Reeve & Walter 1972a; Goto & Yoshida 1997), egg production (Reeve 1970b; Nagasawa 1984), sperm emission (Nagasawa 1987), mating sequence (Reeve & Walter 1972b; Goto & Yoshida 1985; Nagasawa 1985; Goto & Suzuki 2001), fertilization process (Dallot 1968; Ghirardelli 1968; Nagasawa 1985; Goto 1999; Shinn 1994b, 1997), egg laying (John 1933; Dallot 1968; Reeve 1970a, 1970b; Reeve & Lester 1974; Kotori 1975; Goto 1995), germline formation (Carré et al. 2002), establishment of axial body properties (Shimotori & Goto 1999, 2001), neurogenesis (Goto et  al. 1992; Rieger et  al. 2011;

Perez et al. 2013), ultrastructure and ontogeny of the mesoderm-derived tissues and body cavities (Shinn & Roberts 1994; Shinn 1997; Harzsch et al. 2015), and expression of two developmental genes, a median Hox gene (Papillon et al. 2005) and brachyury (Takada et al. 2002).

7.5.1 Mating, fertilization, and egg laying Mating behavior has been mostly studied in benthic species because it is more difficult to keep planktonic ones in laboratory conditions long enough. John (1933), Ghirardelli (1954a, 1968), Duvert et  al. (2000), and Goto and Suzuki (2001) thoroughly described the mating behavior in S. cephaloptera. Chaetognaths are protandrous hermaphroditic animals with only sexual reproduction. First, one mature specimen approaches another and while still attached to the substrate begins to vibrate vigorously the anterior half of its body in a vertical plane. In response, the companion vibrates in turn and approaches the first by two or three short movements. This step is successively repeated several times according to the initial distance of the two mates until they meet side by side. Then they place themselves head-to-tail and suddenly intertwine their bodies before to deposit their spermatophores. In S. cephaloptera, the transfer can be reciprocal or non-reciprocal. The spermatophores are usually deposited dorsally on the neck region, just behind the corona ciliata. Few seconds after the spermatophore has been deposited, the spermatozoa move backward along the dorsal midline of the trunk and separate into two equal masses at the level of the posterior septum before to enter into each opening of the seminal receptacles. Ghirardelli (1968) demonstrated experimentally that when spermatophores are placed on unusual body regions, for instance, on the tail, the migration does not occur normally and spermatozoa are not able to reach efficiently the female genital openings. Duvert et al. (2000) showed that head-less specimens of S. cephaloptera can mate with normal ones, demonstrating that the brain

▸ Fig. 7.42: Location and ultrastructural organization of testes in tail coelom of chaetognaths (Spadella cephaloptera: A–B, E; Ferosagitta hispida: C–D, F–G). Original LM (A) and TEM (B–G) micrographs. A, Transverse semithin (LM) section of tail midregion showing thick bands of primary longitudinal (tail) muscles (tlmu) enclosing two rectangular coelomic cavities (tco), which house numerous cell aggregates at various stages of spermiogenesis (sa). B, Horizontal section of contact zone of both coelomic cavities (with half-cut spermatogenic cells) separated by median extracellular matrix (mecm) secreted at either side by medial (mesenterial) myoepithelial cells and both establishing the tail mesentery. C, Longitudinal section through cluster of spermatogonia (sgoc). D, Some early spermiogenic cells (espc) with typical organelles of the biosynthetic complex. Cross section. E, Section of three different aggregates, two showing cross-cut germ cells in spermatid stage (sptc) displaying electron-dense vesicles (ve) and a flagellar process with clearly visible axonema (ax), a third cluster shows mature spermatozoans (sp) in cross section. F, Several spermatozoans from lumen of seminal vesicle (lsev) in cross-oblique and longitudinal section; note the banded vesicles (bv) overlaying the elongated, extremely elongated nucleus (nu). G, Cross section through posterior nuclear region of several spermatozoans; section taken from proximal oviduct. Further labels: eER, rough endoplasmic reticulum; fc, fin core matrix; lf, lateral fin; go, Golgi stack; mep, multilayered tail epidermis; mi, mitochondrion; ods, oviducal syncytium.

7.5 Reproduction and ontogeny 

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7.5 Reproduction and ontogeny 

is not essential for copulation, at least not for one of the two partners. Goto and Yoshida (1985) found that mating behavior in another member of Spadellidae, Paraspadella gotoi (previously referred as Spadella schizoptera by these authors, see Casanova 1990), is different and more complex than in S. cephaloptera. During the mating sequence, the partners swing their body vertically while still being attached to the substrate and touch each other with their heads. Then while standing up face to face, one mate jumps and transfers its spermatophore to the other. In this species, transfer of spermatophores is thus non-reciprocal and leads to accurate placement of the sperm directly close to the female genital openings. Casanova (1990) referred the unusual ventral position of the female genital openings in contrary to a dorsal positioning in all other chaetognaths and concluded that such a morphological feature is an adaptation to the very peculiar mating behavior of P. gotoi. Regarding planktonic species, mating behavior has only been described in detail for F. hispida (Reeve & Walter 1972b), a chaetognath that could be easily maintained in laboratory culture. Although specimens of this species are usually found in the water column, they exhibit some typical features of benthic chaetognaths such as the ability to attach themselves or their eggs to the substrate, a behavior that may be related to its hyper-neritic distribution (Pierce 1951; Bieri 1991b). Patchy information is available on the mating behavior of another planktonic species, A. crassa (Murakami 1959). Reeve and Walter (1972b) considered the copulation in F. hispida as a variant of cannibalism behavior between two mature specimens of a similar size, a situation in which it is likely much more difficult for one specimen to gain the advantage on the other. These authors also observed that both copulation and cannibalism could occur simultaneously. They suggested that the elaborated mating behavior of S. cephaloptera might have evolved as a mechanism to prevent selfpredation during copulation. In F. hispida, two mature specimens start to swim closely in the water column and make rapid movements for a few seconds. Then one partner grabs the head of its “victim” by means of its

 259

grasping spines. While remaining attached, the two partners engage to swim upward in extreme spiraling motions followed by intervals of rest when they sink downward (Reeve & Walter 1972b). The copulation ends with the spermatophore transfer and the detachment of the mates. Similar to S. cephaloptera, the transfer can be reciprocal or non-reciprocal. The site of attachment is usually on the lateral trunk wall, between the anterior and posterior lateral fins. Immediately after deposition, spermatozoa start to migrate toward the female genital openings such as described in S. cephaloptera. Variation in the mating behavior of planktonic chaetognaths has been reported in A. crassa with mature specimens found adjacent to each other with heads oriented in the opposite direction, presumably copulating (Murakami 1959). In addition to specific mating behaviors, the highly specific morphology of seminal vesicles in some genera like Serratosagitta might serve as copulatory organs whose morphology only fits with the female genital openings of specimens from the same species, preventing hybridization between different species (Ghirardelli 1968). After mating, filiform spermatozoa migrate into the female genital openings (Ghirardelli 1968) and accumulate in the seminal receptacle and the oviducal complex. Then they pass through the oviducal syncytium and the cellular sheath of the oviducal complex in a canal formed by two oviducal specialized cells, the so-called accessory fertilization cells (AFCs), to reach the ovarian space (Stevens 1910; Ghirardelli 1968; Reeve & Lester 1974; Shinn 1992, 1994b, 1997; Goto 1999; Carré et al. 2002) (Fig. 7.46). During ovulation, the zygote moves into the oviducal syncytium through a pore formed by the detachment of degenerative AFCs from the surrounding oviducal cells and enter a membrane-bound lumen in the syncytium (Shinn 1992, 1994b, 1997). Ultrastructural data evidence cytological and functional differences between the two AFCs (Shinn 1994b, 1997). AFC1 forms the extracellular canal allowing the passage of spermatozoa through the oviducal complex prior to fertilization, whereas AFC2 cover the medial side of AFC1 such that only AFC2 is in contact with the

◂ Fig. 7.43: Ultrastructure of tail coelomoduct acting as spermioduct. Ferosagitta hispida. Original TEM micrographs. A, Longitudinal view of left tail nephrostome opening to the tail coelomic cavity (tco), arrow indicates connection between cavity and lumen of sperm duct (spd), nephrostome is lined by numerous multiciliated epithelial cells (mcec). B, Several multiciliated epithelial cells in longitudinal section, three basal bodies (bb) and related ciliary processes (ci) are visible at the apex, lateral portion of plasmalemmae of adjacent epithelial cells intertwine (arrow). C, Detail of most distal part of sperm duct immediately posterior of the nephrostome. D, Distal region of sperm duct with compressed lumen (spdl) lined by distal sperm duct epithelial cells (dsdc). E, Close-up of distal sperm duct cell including a massively developed cytoskeleton of bundled tonofilaments (fib), which are attached to peg-like protuberances of the extracellular matrix (ap); sector indicated by dotted rectangle in D; portion of the tangentially cut, distal sperm duct cells (dsdc*) enclose many electron-dense, presumably resorptive granules (reg). F, Aspect of widened lumen of seminal vesicle (sevl) housing few mature spermatozoans (sp), lumen is surrounded by granulated seminal vesicle-lining epithelial cells (svec). Further labels: bpp, neurites of basiepidermal neuronal plexus (cut longitudinally); ecm, extracellular matrix (of the epidermis); go, Golgi stack; lspc, late spermiogenic cells; ltmu, primary longitudinal (tail) muscles; mi, mitochondrion; miv, microvilli; nu, nucleus; og, highly electron-dense granule; pec, proximal epidermal cells.

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Fig. 7.44: Ultrastructural aspects of proximal region of sperm duct as well as insights from seminal vesicle devoid of deposited sperm. Ferosagitta hispida. TEM original micrographs. A, Cross section of apical part of seminal vesicle with collapsed lumen (sevl), seminal vesicle (duct) epithelial cells (svec) are highly prismatic and tightly packed because of the compression. B, Detail of cross section shown in A (see dotted rectangle) showing numerous electron-dense secretory granules (sg) and further biosynthetic organelles typical for secretory function. C, Proximal part of the (nephridial) sperm duct in oblique section showing widely electron-lucent spermioducal lumen (lsel) surrounded by proximal sperm duct cells (psdc) protruding multiple cilia (ci). D, Close up of apical region of proximal spermioduct cells projecting three cilia into the spermioduct lumen. Further labels: ax, axonema; bb, basal (ciliary) body; cr, cross-striated ciliary rootlet; dec, distal epidermal cells, ecm, extracellular matrix (of the epidermis); fib, bundled tonofilaments running parallel to apical membrane; go, Golgi stack; mi, mitochondrion; nu, nucleus; pec, proximal epidermal cells; rER, rough endoplasmic reticulum.

associated oocyte. The cytoplasm of AFC2 is dominated by vesicles and microtubules and extends in the syncytium with microvilli. Before fertilization, the passage through which the spermatozoa pass is closed because AFC2 has no extracellular canal and its soma obliterate that of AFC1.

The zygote may remain in a syncytium lumen separate from those containing spermatozoa (Shinn 1992, 1997) several hours turned off in meiotic metaphase II as in benthic Spadellidae (Fig. 7.47 B) or release into the seawater 10–15 min after fertilization as in planktonic Sagittidae

7.5 Reproduction and ontogeny 

(Carré et  al. 2002). Completion of meiosis II follows egg laying in all species. Although the occurrence of selffertilization has long been debated (see Ghirardelli 1968), it has been experimentally demonstrated, for instance, by Ghirardelli (1968) in S. cephaloptera, by Dallot (1968) in P. setosa, and by Reeve and Walter (1972b) in F. hispida. However, no data inform whether self-fertilization occurs naturally in the wild and if yes, on its frequency in relation to cross fertilization. Cross fertilization appears to be the usual process as testes and ovaries usually develop consecutively (Kruse 2009), but Shinn (1997) emphasized the weakness of such an argument because spermatogenesis continues during the oogenic period. According to Terazaki and Miller (1982), cross fertilization is the rule even in meso- and bathypelagic species. Planktonic chaetognaths usually release their fertilized eggs directly into the seawater (Doncaster 1902; Dallot 1968; Reeve 1970a, 1970b; Kotori 1975), whereas two benthic Spadellidae, S. cephaloptera (John 1933) and P. gotoi (Goto 1995), and one planktonic Sagittidae, F. hispida (Reeve & Lester 1974), attach clusters of fertilized eggs to the substrate. Immediately after laying, the fertilized eggs of the planktonic species P. elegans are spherical, about 0.3 mm in diameter and suspended in the seawater (Kotori 1975). In P. draco, the fertilized eggs are clumped in a gelatinous masses forming free colonies in the shallow water of about 200–300 eggs each (Shimotori et  al. 1997). They hatch within the gelatinous mass, and juvenile are released into the seawater after four days. In larger specimens of S. cephaloptera, up to eight eggs are discharged at the same time from each ovary and attached to the substrate (Fig.  7.47  C; see also John 1933). Several deep sea-inhabiting Eukrohnia species, such as E. hamata (Dawson 1968), E. bathyantarctica (Alvariño 1968), E. fowleri (Terazaki & Miller 1982), and E. bathypelagica (Terazaki & Miller 1982), broods the eggs internally in kind of marsupial sacs (Fig. 7.47 F–I).

7.5.2 Germline, cleavage, gastrulation, and coelomogensis Elpatievsky (1909) was the first to report that the germline was derived from a blastomere that contained a “cytoplasmic body” near the vegetal pole of the zygotes as detectable shortly before the first mitosis. More recently, the fate of the primordial germ cells (PGCs) has been thoroughly studied in several species by Carré et  al. (2002). After spawning, circular movements aggregate cytoplasmic material located in the vegetal cortex of the fertilized eggs and lead to a single germ granule, the autonomous

 261

formation of which is linked to meiotic and mitotic cell cycle factors and does not depend on factors brought by spermatozoa. The germ granule is segregated into one of the two first blastomeres and is inherited in a unique vegetal blastomere until the 32-cell stage. At the 32-cell stage, the germ granule segregates into the smallest blastomere constituting the founder PGC. Then the germ granule is partitioned into two presumptive PGCs further divided into four PGCs that become the female and the male germlines in hatchling. Immunohistochemistry demonstrated the presence of a vasa-like protein within the germ granule. The presence of a vasa-like protein has been demonstrated in the germ granule, in PGCs, and in cytoplasmic material associated with the germinal vesicle of oocytes of chaetognaths as in many other metazoans such as Nematostella, Tubifex, Platynereis, Daphnia, Tetranychus, Parhyale, Drosophila, and Medaka (reviewed in Özhan-Kizil et al. 2009). So far, the function of vasa-like protein is unknown in chaetognaths. The embryonic cell cleavage is total and equal (reviewed in Brusca & Brusca 2003), except for one unequal cleavage, which segregates the germ granule into the founder germline cell at 32-cell stage (Carré et al. 2002). In many studies, the early development of chaetognaths has been repeatedly described as radial and similar to deuterostomes (reviewed in Harzsch et al. 2015). However, recent data based on injections with fluorescent lineage tracer in single blastomere at 2- and 4-cell stages lead new insights on chaetognath relationships (Shimotori & Goto 1999, 2001) and corroborated previous observations from Elpatievsky (1909). These authors reported after the second mitosis a shift in a counterclockwise (leiotropic) displacement of the two animal cross-furrow cells with respect to the two vegetal cross-furrow cells, the next cleavage occurring in the opposite direction. Such an early embryo cleavage is also seen in spiralian protostomes. Shimotori and Goto (2001) also emphasized the link between the future body axes and the tetrahedral disposal of the blastomeres at the 4-cell stage, a situation also resembles that of spiralian protostomes. These authors concluded that contrary to the classical views, the early embryo cleavage of chaetognaths displayed more similarities with protostomes than with deuterostomes. Our knowledge of their gastrulation, mesoderm formation, and coelomogenesis comes from the classical accounts of Hertwig (1880), Doncaster (1902), Burfield (1927), John (1933), Shinn (1994a), and Shinn and Roberts (1994) on several species belonging to Spadellidae and Sagittidae, and has been reviewed by Shinn (1997) and Harzsch et  al. (2015) (Fig.  7.48). The entoderm forms by invagination of the blastopore. Although the topic has been

262 

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Fig. 7.45: Ovarian system in the trunk of chaetognaths including bilaterally symmetrical ovaries and the oviducal complex. A–B, Transverse semithin (LM) sections of the most posterior region of the trunk (A) and at the transition zone of trunk and tail (B) of Spadella cephaloptera. Pair of transverse (phragma) muscles (trmu) is marked in A, phragma muscles are crammed in interspace built mediany by lateral faces of intestine (int) and ovaries (ov) as well as laterally by primary longitudinal (trunk) musculature (tlmu). B. Posterior (proximal) end of the oviducal complex (gd) passing into the vagina (va) and opening outside to side of the body via the female gonopore (fgp); position of secondary (homosarcomeric) muscles is indicated by arrows. Originals. C–F: Original TEM micrographs. Ferosagitta hispida. C–D: Ultrastructural details of ovarian contents. C, Previtellogenic oocyte (ocp) charactericed by thin oolemma (oo) and polymorphic, highly

7.5 Reproduction and ontogeny 

 263

Fig. 7.46: Fertilization process in Paraspadella gotoi. A, Fully grown oocyte associates with two accessory fertilization cells (AFCs). B, AFC2 sinks in the oocyte and occludes the fertilization extracellular canal formed in AFC1. C, The cytoplasmic process of AFC2 disappears from the fertilization canal through which filiform spermatozoid invades the egg. D, A single spermatozoid invades an oocyte after germinal vesicle breakdown (GVBD). AFC2 moves outside from the oocyte. E, Fertilized oocyte moves into the syncytium oviducal complex through a pore that is formed by degeneration of AFCs (DeAFCs). Spermatozoid chromatin is condensed as a round shape. F, Fertilized oocyte is stored in the oviducal complex at the first meiotic metaphase until being laid in sea water. Resumption of meiosis occurs after being laid in sea water. DeAFCs, degenerating AFCs; OC, oviducal complex; OS, oviducal syncytium; OW, oviducal cellular wall; Sp, sperm. (Redrawn and modified from Goto 1999, including ultrastructural descriptions of Shinn 1992, 1994b, 1997.)

a subject of controversy, chaetognaths are definitely true triploblastic bilaterians (Duvert & Salat 1979; Ahnelt 1980; Welsch & Storch 1982; Shinn 1994a; Shinn & Roberts 1994; reviewed by Shinn 1997 and Harzsch et al. 2015), but as in the case of the early cleavage, which has been erroneously considered radial, the fate of the blastopore and the ontogeny of the three germ layers of the embryo (gastrulation and coelomogenesis) have been mistaken. Specifically, the endo-mesoderm formation by folding the entoderm and the secondary formation of the mouth at the opposite of the blastoporal site have been used as the main developmental features to bring together chaetognaths and deuterostomes (Kuhl 1938; Hyman 1959). However, Kapp (2000) discouraged the use of the term “enterocoely”. She noticed that although gastrulation and mesoderm formation in chaetognaths conform to some aspects of the deuterostome developmental pattern, they also display important variations. Indeed, in the typical enterocoely, as found in lower deuterostomes such as Ambulacraria, the mesoderm arises by folding of the entoderm toward the

outside of the embryo (outpouchings of the archenteric cavity), which form the coelomic cavities after their complete separation from the entoderm. On the contrary, in chaetognaths, the mesoderm forms by entoderm folding, which progresses backward directly into the archenteron (inpouchings) toward the blastoporal site. The enterocoelic nature of the body cavities in chaetognaths has been also questioned by Meglitsch and Schram (1991) who considered their coelom not homologous to archimeric lower deuterostomes. This peculiar mode of early coelomogenesis has been termed “heterocoely” by Kapp (2000), who emphasized the difficulties to relate these developmental features to other bilaterians. Indeed, the formation of definite bilateral coelomic cavities in trunk and tail could be interpreted as a variant of schizocoely, which implies the formation of coelomic cavities from compact masses of mesodermal cells which transform into a coelothel then surrounding a coelomic cavity (Schmidt-Rhaesa 2007). The trunk coelom derives from bases of primary longitudinal muscle cells

◂ electron-dense spots (nlm) abutting the central nucleus (nu). D, Follicular reticulum (fr) diversifying through ovarian space, locally restricted to small to small compartments (osc); note the osmiophilic cortical granules (cg) typical for vitellogenic oocytes (ocv). E–F: Ultrastructural details of the oviducal complex. E, Oviducal canal cell (odc) in cross section, cytoplasm is rich in swollen cisternae of the rough endoplasmic reticulum (rER). F, Oviduct in longitudinal section, aggregation of spermatozoans (sp) is encompassed by oviducal syncytium (ods) and oviducal canal cells (odc), tightly bundled myofilaments (my) run parallel to the apical membrane of oviducal canals cells. Further labels: cto, ciliary tuft organ; eg, fertilized egg; mep, multilayered epidermis; mi, mitochondria; miv, microvilli; tco; coelomic space of the trunk; vgp, ventral gland papillae.

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Fig. 7.47: Some developmental aspects in chaetognaths. A–E, Original light micrographs. Spadella cephaloptera. A, Trunk region of a living specimen with fully grown oocytes (oo) in the ovaries. B, Fertilized and ovulated oocytes stored in the oviducal complex at the first meiotic metaphase until being released into the sea water. Note their polyhedral shape and the nuclear membrane disappearance after germinal vesicle breakdown (GVBD) when compared with fully grown oocytes. C, Laying eggs with early embryos attached in cluster to the surface of a glass container. D, E, Newly hatched young. h, head; tr, trunk; vnc, ventral nerve center. F–I, light micrographs after Kruse (2010, http:// hdl.handle.net/10013/epic.34655). E. bathyantarctica. Posterior part of the trunk at different stages of development. F, Ovaries (ov) with oocytes and the female genital opening (arrow). G, H, Developing juveniles in marsupial sacs. In Eukrohnia species, the lateral fins form a brood pouch to cover the developing eggs. The juveniles are retained in these marsupial sacs for at least some time after hatching. I, Empty marsupial sacs.

successively drifting apart to form compact muscle pads in the dorsal or ventral rim of the trunk. Incipient coelomic spaces then merge in the process of drifting and final differentiation of the primary longitudinal musculature (Shinn & Roberts 1994; Shinn 1997). The periovarian coelom becomes established when the female germ cell proliferates and loosens from precursors of also proliferating circumintestinal myoepithelial cells and lateral field cells. The coelomic cavity becomes continuously larger because the surrounding myoepithelial cells secrete coelomic liquids into the lumen. The same mechanism of coelomogenesis leads to the formation of both coelomic spaces in the tail (Shinn & Roberts 1994; Shinn 1997) Thus, the formation of peri-ovarian and tail coeloms can also be assumed to work in a similar way

as documented in protostomian Bilateria performing schizocoely. This striking mixture of coelomogenetic patterns, either autapomorphic or shared with deuterostomes or protostomes, motivated a study of the expression of the brachyury gene, a key factor for the mesoderm formation, during the early and late development of P. gotoi (Takada et al. 2002). The expression of brachyury was found specifically not only at the blastopore site and at the opposite of the early embryo as in basal deuterostomes but also around the mouth opening region of the late embryo. Thus, their gastrulation and coelomogenesis are clearly distinct from others protostomes and cannot be used as derived shared features to consider them as in-group deuterostomes (Perez et al. 2014; Harzsch et al. 2015).

7.5 Reproduction and ontogeny 

7.5.3 Hatching and growth Duration from egg laying to hatching depends on the species considered and breeding conditions. It is about 24 h at 21–24°C in S. cephaloptera (Y. Perez & C.H.G. Müller, personal observations), 24 h at 16–18°C in A. crassa (Murakami 1959), and 30 h at 23°C in P. draco (Shimotori et al. 1997). In P. gotoi, it is about 2 days at 23°C and reaches 4 days at 17°C (Shimotori et  al. 1997). Chaetognaths are direct developers and become sexually mature in about 1–2 months (Reeve 1970a, Reeve & Walter 1972a, Reeve & Lester 1974). The architecture of mesodermal tissues in the hatchling and juvenile has been thoroughly studied by Shinn and Roberts (1994) and Shinn (1994a, 1997) in F.  hispida, and recently by Harzsch et  al. (2015) in S. cephaloptera. In the latter species, hatchling displays two body segments, the head and trunk, separated by the anterior septum which forms the boundary between the cephalic and trunk mesoderm (Harzsch et  al. 2015). F. hispida hatches in a more advanced state of differentiation, and the rudiment of the posterior septum is already formed by specialized peritoneocytes, which enfold the PGCs (Shinn & Roberts 1994; Shinn 1994a, 1997). Moreover, numerous small triangular spaces appear in the developing mesoderm of F. hispida (Shinn & Roberts 1994; Shinn 1994a, 1997), whereas it is compact without any trace of cavitation in S. cephaloptera (Harzsch et al. 2015). These differences could rely on species-specific variations in the tissue level differentiation or on different experimental breeding conditions (Harzsch et al. 2015). It has been postulated that the coelomic cavities arise from the coalescence of these lacunae (Shinn & Roberts 1994; Shinn 1994a, 1997). The full adult morphology develops several days after hatching according to the species considered (John 1933; Goto & Yoshida 1985; Shinn & Roberts 1994; Shinn 1994a, 1997; Harzsch et al. 2015). In S. cephaloptera, stereotypically arranged coelothelial cells form a continuous wall that delimits trunk and tail cavities 2–3 days after hatching. The cephalic chitinous structures, the gut, the brain, the eyes, and the corona ciliata are also complete. According to Harzsch et al. (2015), the late differentiation and cavitation of the coelothel from two compact mesodermal bands as found in S. cephaloptera may be regarded as a derived, postembryogenic variant of the schizocoely found in some annelids. These ultrastructural data strongly suggest that chae­ tognaths should be considered fundamentally dimeric bilaterians that become secondarily trimeric after the completion of the posterior septum (Harzsch et al. 2015), which coincides with the definitive segregation of the

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female and male germlines, in the trunk and tail segments, as previously proposed by Kowalevsky (1871), Doncaster (1902), John (1933), and Kapp (2000). The head, trunk, and tail coelomic cavities in adult chaetognaths are not homologous to those of archimeric animals and cannot be used to relate chaetognaths with deuterostomes (Meglitsch & Schram 1991; Harzsch et al. 2015). Given these facts and according to the latest genomic data inferring their pivotal phylogenetic position as basal protostomes, the embryonic development of chaetognaths, although autapomorphic in many aspects, could represent an intermediate evolutionary step to modify the typical enterocoely of some deuterostomes toward the derived schizocoely of spiralian protostomes (Harzsch et al. 2015).

7.5.4 Neurogenesis At hatching, the cephalic nervous system is represented by a simple fiber loop that extends anteriorly from the ventral nerve center. This “primary brain” surrounds the area where the secondary mouth will open. It is topologically a circumoral nerve ring (Rieger et  al. 2011) and will develop into the posterior brain domain of the adult. This “primary sensory brain” will receive input from the sensory organs a few days later and will be involved in the modulation of motor behaviors in response to changing sensory input (Rieger et al. 2010, 2011). During the first 48 h after hatching, synaptic zones emerge in the brain neuropil, new RFamide-like immunoreactive neurons appear, and a system of RFamide-like immunoreactive fibers is elaborated, all of which implies that a functional brain is present by the time the hatchlings switch from feeding on yolk supplies to active predation. Postembryonic neurogenesis also includes the emergence of the second brain component that topologically can also be viewed as being arranged in a circumoral pattern (Rieger et al. 2011). This anterior brain domain will give rise to the circumesophageal and stomatogastric nervous system, which, in the adult, is involved in controlling the grasping spines as well as the musculature activity responsible for opening and closing the mouth (Rieger et al. 2010). Similar to the situation in the adult, the developing ventral nerve center most likely modulates body movements in hatchlings, enabling them to escape predators and control their attachment to seaweed, the preferred substrate for attachment in this species. A comparison of the system of neurons that express RFamide‐related neuropeptides showed that the hatchling pattern in many respects resembles that in the adult. The number of somata with RFamide‐like immunoreactivity increases very little from hatching onward,

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Fig. 7.48: The developmental sequence in Sagitta bipunctata and Spadella cephaloptera. A–D, Early cleavage in S. bipunctata (after Elpatievsky 1909); E–L, early cleavage in S. cephaloptera; M–O, gastrulation and mesoderm formation in S. cephaloptera; P, specimen of S. cephaloptera just before hatching; Q–S, gastrulation and mesoderm formation in S. bipunctata (after Burfield 1927). A few hours after the beginning of the cleavage, the blastopore (bl) completely closes. Two mesodermal folds progress backward directly into the archenteron (Ar) and mark off the endoderm (end) from a pair of mesodermal cavities (hc, head coelom; tc, trunk coelom). The anterior region of the

Literature 

but the system of their neurites becomes more complex as development proceeds (Harzsch et  al. 2009; Rieger et al. 2011). Labeling of dividing neuronal progenitors with the s-phase-specific mitosis marker bromodeoxyuridine (BrdU) evidenced extensive level of mitotic activity in the ventral nerve center and the brain for ca. 3 days after hatching (Perez et  al. 2013). Neuronal progenitors cycle rapidly, likely divide in an asymmetric fashion and are arranged in a distinct grid‐like geometrical pattern, including about 35 transverse rows of about five to six cells. This two-dimensional arrangement is not optimal and forms a triangular point system so that each progenitor is in contact with four others. This implies that possible biochemical constraints related to cell-to-cell interactions and mitotic spindle orientation may define the highly geometric arrangement of neural progenitors in chaetognaths (Perez et  al. 2013). Moreover, their iterated geometrical pattern may be interpreted as revealing serially organized domains of the developing ventral nerve center as it has been previously proposed in the adult on the basis of serially iterated RFamide-like immunoreactive neurons (Harzsch et  al. 2009). BrdU labeling experiments also showed mitotic activity in the developing corona ciliata and within developing epidermal ciliary sense organs. Interestingly and as previously observed in the corona ciliata by Shinn (1997) on the basis of ultrastructural data, the mitotic activity persists after the postembryonic development attesting to a turnover of sensory neurons during adult life. The expression of the Hox gene SceMed4 (a putative ortholog to Antp or Scr group) has been analyzed in the late embryo, hatchling, and juvenile of S. cephaloptera (Papillon et  al. 2005). SceMed4 expression is restricted in the ventral nerve center in a typical position-specific pattern. This suggests that Hox genes may play primarily a potential role to generate the neuronal diversity and to regionalize the central nervous system during neurogenesis (Papillon et al. 2005). Recently, the analysis of Hox gene expression during embryogenesis of the rotifer Brachionus manjavacas also showed a restricted expression pattern restricted in the central nervous system (Fröbius & Funch 2017). In conclusion, the analysis of chaetognath neurogenesis and comparative expression of developmental genes

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in the central nervous system provided evidence that (1) a serially repeated organization in the central nervous system may be one the first steps toward true metamerism found in higher bilaterians such annelids, arthropods, and chordates, and (2) Hox genes may primarily play a potential role to generate the neuronal diversity and regionalize the central nervous system rather than the usual expression patterns along the anterior-posterior axis.

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◂ ectoderm (ect) forms the stomodeum (st). T, Specimen of S. bipunctata just before hatching (modified from Doncaster 1902). Early cleavage is total and equal. Labels denote the 2-cell (2c), 4-cell (4c), 8-cell (8c), etc., stages and the developmental time in minutes and hours from fertilization. Ar, archenteron; bl, blastopore; d, descendants of the d cell; ect, ectoderm; ent, entoderm; h, head; hc, head coelom; mes, mesoderm; PGCs, primordial germ cells; RK, “Richtungskörper” = polar body; st, stomodeum; tc, trunk coelom, vnc, ventral nerve center; X, germ granule (reprinted with permission from Harzsch et al. 2015).

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Caecosagitta macrocephala, With Further Observations on Bioluminescence in Chaetognaths. Biological Bulletin 219: 100–111. Tokioka, T. (1939): Three new chaetognaths from Japanese waters. Memoirs of Imperial Marine Observatory, Kobe Marine Observatory 7: 129–140. Tokioka, T. (1940): A small collection of chaetognaths from the coast of New South Wales. Records of the Australian Museum 20: 367–379. Tokioka, T. (1956): On chaetognaths and appendicularians collected in the central part of the Indian Ocean. Publications of the Seto Marine Biological Laboratory 5: 197–202. Tokioka, T. (1959): Observations on the taxonomy and distribution of chaetognaths of the north Pacific. Publications of the Seto Marine Biological Laboratory 7: 359–456. Tokioka, T. (1965a): The taxonomical outline of chaetognaths. Publications of the Seto Marine Biological Laboratory 12: 335–357. Tokioka, T. (1965b): Supplementary notes on the systematics of Chaetognatha. Publications of the Seto Marine Biological Laboratory 13: 231–242. Tokioka, T. (1974a): Morphological differences observed between the generations of the same chaetognath population. Publications of the Seto Marine Biological Laboratory 21: 269–279. Tokioka, T. (1974b): On the specific validity in species pairs or trios of plankton animals, distributed respectively in different but adjoining water masses, as seen in chaetognaths. Publications of the Seto Marine Biological Laboratory 21: 393–408. Tönnesson, K. & Tiselius, P. (2005): Diet of the chaetognaths Sagitta setosa and S. elegans in relation to prey abundance and vertical distribution. Marine Ecology Progress Series 289: 177–190. Tse, P., Hui, S.Y. & Wong, C.K. (2007): Species composition and seasonal abundance of Chaetognatha in the subtropical coastal waters of Hong Kong. Estuarine Coastal and Shelf Science 73: 290–298. Ulloa, R., Palma, S. & Silva, N. (2000): Bathymetric distribution of chaetognaths and their association with water masses off the coast of Valparaiso, Chile. Deep Sea Research Part I: Oceanographic Research Papers 47: 2009–2027. Van der Spoel, S., Heyman, R.P. (1983): A Comparative Atlas of Zooplankton: Biological Patterns in the Oceans. Springer-Verlag, Berlin. Vannier, J. & Chen, J. (2005): Early Cambrian food chain: new evidence from fossil aggregates in the Maotianshan Shale biota, SW China. Palaios 20: 3–26. Vannier, J., Steiner, M., Renvoisé, E., Hu, S.X. & Casanova, J.P. (2007): Early Cambrian origin of modern food webs: evidence from predator arrow worms. Proceedings of the Royal Society B, Biological Sciences 274: 627–633.

Villenas, F. & Palma, S. (2006): Sagitta chilensis nueva especie de quetognato en fiordos australes chilenos (Chaetognatha, Aphragmophora, Sagittidae). Investigaciones marinas, Valparaíso 34: 101–108. Vinogradov, M.E. (1997): Some problems of vertical distribution of meso-and macroplankton in the ocean. Advances in Marine Biology 32: 1–92. Vega-Pérez, L.A. & Schinke, K.P. (2011): Checklist of Chaetognatha phylum from São Paulo State, Brazil. Biota Neotropropica 11: 541–550. Wada, H. & Satoh, N. (1994): Details of the evolutionary history from invertebrates to vertebrates, as deduced from the sequences of 18S rDNA. Proceedings of the National Academy of Sciences, USA 91: 1801–1804. Walz, B. (1974): The fine structure of somatic muscles of Tardigrada. Cell Tissue Research 149: 81–89. Walz, B. (1975): Ultrastructure of muscle cells in Macrobiotus hufelandi. Memorie dell’Istituto Italiano di Idrobiologia 32 (Suppl.): 425–443. Welsch, U. & Storch, V. (1982): Fine structure of the coelomic epithelium of Sagitta elegans (Chaetognatha). Zoomorphology 100: 217–222. Welsch, U. & Storch, V. (1983a): Fine structural and histochemical observations on the epidermis and the sensory cells of Sagitta elegans (Chaetognatha). Zoologischer Anzeiger 210: 34–43. Welsch, U. & Storch, V. (1983b): Enzymehistochemische und elektronmikroskopische Beobachtungen am darmepithel von Sagitta elegans. Zoologische Jahrbücker, Abteilung für Anatomie 109: 23–33. Wei, S., Li, P., Yang, M., Zhou, L., Yu, Y., Ni, S., Wang, X. & Qin, Q. (2016): The mitochondrial genome of the pelagic chaetognath, Pterosagitta draco. Mitochondrial DNA Part B 1: 515–516. Willmer, P. (1990): Invertebrate relationships: Patterns in Animal Evolution. Cambridge University Press, Cambridge. Winkelmann, C., Gasmi, S., Gretschel, G., Müller, C.H.G. & Perez, Y. (2013): Description of Spadella valsalinae sp. nov., a neo-endemic benthic chaetognath from Northern Adriatic Sea (Croatia) with remarks on its morphology, phylogeny and biogeography. Organism Diversity & Evolution 13: 189–202. Wu, X., Li, K., Huang, L. & Yin, J. (2014): Seasonal and spatial distribution of chaetognaths on the north-west continental shelf of the South China Sea. Journal of the Marine Biological Association of the United Kingdom 94: 837–846. Zink, R.M. & Barrowclough, G.F. (2008): Mitochondrial DNA under siege in avian phylogeography. Molecular Ecology 17: 2107–2121. Zrzavý, J., Mihulka, S., Kepka, P., Bezděk, A. & Tietz, D. (1998): Phylogeny of the Metazoa based on morphological and 18S ribosomal DNA evidence. Cladistics 14: 249–285.

Kenneth M. Halanych, Michael G. Tassia and Johanna T. Cannon

8 Pterobranchia 8.1 Introduction

Pterobranchs are small suspension-feeding colonial hemichordate deuterostomes that live in marine environments within a secreted tubular structure called a coenecium (Figs. 8.1 and 8.2). Individual zooids have a feeding apparatus with paired tentaculate arms that emerge from the middle region of their tripartite bodies. Extant pterobranchs have limited diversity with 23 recognized species in two genera, Cephalodiscus M’Intosh 1882 and Rhabdopleura Allman 1869b (Fig. 8.1), which together are the sister taxon to acorn worms also callled enteropneust hemichordates (Tassia et al. 2016; Cannon et al. 2014). G.O. Sars collected pterobranchs in 1866 in northern Norway, but Allman was the first to formally describe the species, Rhabdopleura normani, in 1869a, b. Cephalodiscus was collected a short time later, 1876, on the H.M.S. Challenger expedition in the Strait of Magellan and described by M’Intosh (1882). Although Sars is often credited as the first to collect pterobranchs, Cephalodiscus was collected earlier, in either 1841 or 1842, during the Erebus and Terror Expedition to the Antarctic—a fact not recognized until sometime between 1883 and 1887 (Ridewood 1912, 1921). Early workers allied these novel animals with bryozoans, hydroids, and other “polyzoa”, but Lankester (1877) coined the term Pterobranchia (Greek—Pteron, wing or feather; Old French—branche, gill) and placed Rhabdopluera within Bryozoa. In 1887, two workers revisited pterobranch affinities. M’Intosh (1887) places both genera within Bryozoa, whereas Harmer (1887) formally recognized that Cephalodiscus, which has a pair of gill pores, was related to acorn worms and proposed moving Cephalodiscus into Hemichordata. The current classification of hemichordates recognizing Pterobranchia as one of two classes within Hemichordata was proposed by Willey (1899a, b). For additional details on the history of Hemichordata, see Chapter 9 on Enteropneusta, this volume. In addition to modern species, a close association between Rhabdopleura and graptolite fossils was recognized almost immediately (Allman 1872) and has been solidified by recent works (e.g., Crowther & Rickards 1977; Mitchell et al. 2013). Thus, pterobranchs have a fossil history spanning back to the middle Cambrian. Because of the architecture of coenecia in pterobranchs, graptolites are considered to be more closely related to https://doi.org/10.1515/9783110489279-008

Rhabdopleura than Cephalodiscus. Although graptolites have been well studied, herein we focus on extant forms, which have a very limited literature. Pterobranchs are small, and larger colonies live below 100 m, limiting them as subjects of biological studies. Hyman’s (1959) treatment of pterobranchs, in many ways, still represents current knowledge and is arguably one of the most complete pterobranch treatments in the last 80 years (but see Benito & Pardos 1997).

8.2 Morphology 8.2.1 General anatomy Adult pterobranch individuals, or zooids, are typically 1–6 mm in length (Halanych et al. 2012) but have been reported up to 14 mm (John 1931). The colonial structures they live in, however, can be 20–30 cm in some species of Cephalodiscus. All currently recognized species of extant pterobranchs secrete a coenecium, which is a tubular structure that protects the colony. Rhabdopleura compartmentalizes individual zooids within typically dark-brown to black coenecia, and individual zooids are connected by a dark organic stolon. Branching of the stolon typically follows a bifurcating pattern, but this has not been well examined in several species. This construction plan of coenecia typically results in encrusting colonies or, at least, colonies that do not rise far above the substrate in extant species (Fig. 8.2 D). By contrast, some graptolites used a similar plan to form large colonies. All species of Rhabdopleura have one pair of tentaculate arms that emanates from there collar region, and they lack external gill pores. Comparatively, Cephalodiscus species do not compartmentalize zooids within the coenecium to the same degree. Whereas some species (e.g., Cephalodiscus fumosus John 1931) have zooids more or less constrained to individual tubes, hundreds of zooids can occupy a common coenecial space in other taxa (e.g., Cephalodiscus hodgsoni Ridewood 1907b). Cephalodiscus includes encrusting species (e.g., Cephalodiscus gracilis Harmer 1905) as well as several species (C. hodgsoni, Cephalodiscus nigrescens Lankester 1905, and Cephalodiscus densus Andersson 1907) which build large structures that are

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a

md gs

co

cs an

cs

tr

stl

elevated from the benthos allowing zooids to get out of the benthic boundary layer to aid suspension feeding. Most Cephalodiscus zooids range from a bright orange to deep red to brown color, and the coenecium is often a translucent yellowish to orange color. Some species, such as C. fumosus and C. densus, can agglutinate large amounts of sediment to the tube obscuring the tube altogether. For Cephalodiscus species that have shared coenecia, individuals form clusters of zooids related by asexual reproduction that vary in developmental age (Fig. 8.2 C). These clusters have been observed to range in size from a single individual to groups of about 20 zooids (Halanych, personal observation). Larger clusters may have multiple mature and reproductively active zooids. Adult Cephalodiscus zooids have three to nine pairs of tentaculate arms and have a pair of pharyngeal gill pores. These openings are often referred to as gill pores because they are more rounded than slit-like in shape. Individual zooids are divided into three body regions that correspond to internal coelomic partitions (see Section 8.2.3, “Body Cavities”; Fig. 8.3). They are referred to as the cephalic shield, collar (with feeding arms), and trunk (with a posterior stolon). The anteriormost region of the body, or cephalic shield, is a broad rounded structure that acts much like a molluscan foot. The ventral surface of the cephalic shield is densely ciliated, and zooids can use it to glide across the substrate. The ventral surface of the cephalic shield contains a distinct band that is

co

Fig. 8.1: Generalized body plans of pterobranchs. Cephalodiscus in on the left and Rhabdopleura on the right. Note that Rhabdopleura does not usually extend this far out of the tube. a, arms with tentacles; an, anus; co, collar; cs, cephalic shield; gp, gill pore; md, mesocoel duct; stl, stolon; tr, trunk. Illustration A from Lester et al 1985.

associated with a change in ciliation pattern (Halanych, personal observation), and in Cephalodiscus this region is also often visible as a bright red pigmented strip (Hyman 1959; Benito & Pardos 1997). The significance of the pigmented strip is not known. In both genera, the cephalic shield is also the primary organ used in tube building. The cephalic shield secretes a mucous-like substance from ventrally oriented secretory cells to form the coenecium. The behavior of tube building has not been well studied, but both Cephalodiscus and Rhabdopleura zooids will cup their cephalic shield along the aperture of the coenecium. This pose is likely done to aid tube building, as well as to permit extension of the arms out of the coenecium while feeding. If removed from the coenecium, pterobranchs have the ability to build a coenecium de novo. The collar region possesses the mouth and pairs of tentaculate feeding arms that are invaded by a mesocoelomic cavity and a blood vessel. In Rhabdopleura, the two arms straddle the dorsal medial line. Cephalodiscus, with multiple pairs of arms, has a similar arrangement, but the arms form a crescent pattern with the youngest arms most posterior and slightly more lateral. The overall shape, function, and morphology of these arms fits the classical definition of a lophophore (Halanych 1996), but that term has typically not been associated with this group of organisms although earlier workers recognized the similarity (Allman 1869a, b; Hyman 1959; Gilmour 1979; Hoverd 1985; Lester 1985; Dilly et al. 1986c). As in acorn

8.2 Morphology 

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Fig. 8.2: Representatives of live pterobranchs. A, Bermudan Cephalodiscus gracilis zooid in feeding position. B, Bermudan Rhabdopleura normani zooid in feeding position. C, Coenecium of Cephalodiscus hodgsoni. D. Coenecium of Rhabdopleura normani. Scale bars in lower right for A, B, and D are 1 mm. Scale bar for C is 1 cm.

worms, the collar region is heavily involved in manipulating and controlling which food particles enter the mouth. The most posterior region is the trunk, which hosts the major internal organs. In contrast to the cephalic shield and collar region, the trunk is more sparely ciliated externally. Just below the collar, on the anterior trunk region, Cephalodiscus possess a single pair of gill pores that are not found in Rhabdopleura. Due to life in a tube, pterobranchs possess a U-shaped digestive track with an anus exiting dorsally near the anterior end of the trunk. A single gonad, or pair of gonads, is also found in the trunk regions with genital pores opening on either side near the anus. The posterior region of the trunk tapers into a long muscular stolon, which can extend several millimeters exceeding the length of the zooid. In the case of Rhabdopleura, this stolon connects to other

zooids in a bifurcating pattern, whereas in Cephalodiscus, the stolon terminates in a pedal disk that is a proliferation zone for asexually derived zooids. This pedal disk also serves as a point of non-permanent attachment for the cluster to the substrate. The glandular epithelium in the pedal disk (Hyman 1959), and the occasional difficulty of removing the disk from the substrate, suggests that the attachment to the substrate is more than muscular and secretions are likely involved. Drawings in earlier publications often show single zooids with the stolon just cut off at the end and lacking a pedal disk (e.g., Hyman 1959, figs. 56 B and 63 A). These zooids were presumably drawn after they had been plucked from a cluster, as termination of the stolon in living Cephalodiscus is not shaped like this (see Hyman 1959, fig. 58 A with “glandular, center of stalk end”).

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Fig. 8.3: Representation of the internal morphology of a Rhabdopleura zooid. Position of gill pores in Cephalodiscus is indicated. a, arms with tentacles; an, anus; cg, collar ganglion; co, collar; cs, cephalic shield; csm, cephalic shield musculature; dv, dorsal blood vessel; e, esophagus; g, glomerulus; gd, gonad; gp, gill pore; h, heart; i, intestine; mc, mesocoel; md, mesocoel duct; mo, mouth; p, pericardium; pc, protocoel; pp, protocoel pore; s, stomach; stl, stolon; tc, trunk coelom; tr, trunk. Illustration from Benito and Pardos (1997).

The relative orientation of the adult body plan deserves note. Herein we consider the cephalic shield to be anterior with the stolon posterior and the tentaculate arms to be dorsal with the ciliated surface of the cephalic shield to be ventral. Hyman (1959) questions this interpretation based on descriptions of development by earlier workers (who she does not specify but most likely Gilchrist 1917 and John 1932). Instead, similar to a phoronid or tunicate, which also possess U-shaped guts, Hyman considered most of the trunk’s surface to be ventral with the short region between the mouth and the anus to constitute the dorsal surface. In other words, she strictly interpreted body orientation based on primary body axis as defined by the mouth and the anus.

8.2.2 Integument Characterization of the pterobranch integument has been limited, but see Andersson (1907) and Welsch (1984). The epidermis consists of a squamous epithelium with a single

layer of cells, and the basement membrane under the epithelium is similar to that found in acorn worms. Most cells are monociliated with cilia containing a typical 9 × 2 + 2 microtubule pattern, and they cover most of the zooid, although the density of the cilia varies considerably. Glandular cells, that are presumed to be mostly mucous producing, are present throughout. The epithelium of the pedal disc in Cephalodiscus also possesses a thickened epithelium (Hyman 1959, fig. 58) and presumably some secretory cells which aid in the attachment of the cluster to the substrate. The most complex epithelium is found on the ventral surface of the cephalic shield, which is heavily ciliated. One of the main functions of the cephalic shield is to secrete the coenecium and thus, the ventral surface possesses regions of thickened epithelium with taller cells whose main function is secretory. Based on cellular complexity and given the presence of a prismatic basic cell type and four types of glandular cells, Welsch (1984) comments on the similarity between the ventral surface of the cephalic shield and the proboscis of acorn worms. In addition, the ventral surface contains the pigmented

8.2 Morphology 

strip, which is about three to four cells wide, and contains osmiophilic granules (Benito & Pardos 1997).

8.2.3 Body cavities Pterobranchs are tripartite in nature with an unpaired protocoel, a paired set of mesocoels, and a paired set of metacoels. Studies examining the development of pterobranchs are limited (see Section 8.3, “Reproduction and Development”), and thus not much is known about the formation and ontogeny of coelomic cavities in these animals. The protocoel is unpaired and delineated from the mesocoel by a septum. Hyman (1959) reports that the protocoel only extends posteriorly to the pigmented stripe in Cephalodiscus. The heart vesicle projects anteriorly, and its presence creates two dorsolateral pockets of the protocoel. These pockets are heavily ciliated and lead into narrow tubes that exit the body through protocoel pores. A pericardial cavity forms a large sinus in the protocoel (see Section 8.2.7, “Excretory and Circulatory Systems”). The paired mesocoels surround the buccal region and abut each other by forming dorsal and ventral septa. In addition, mesocoelomic cavities invade the feeding arms and tentacles and act as a hydrostatic skeleton for these appendages. As mentioned above, the mesocoelinvaded feeding apparatus in pterobranchs fits the classic definition of a lophophore (Gilmour 1979; Hoverd 1985; Halanych 1996). A pair of mesocoel extensions also reach into the protocoel and are associated with the heart vesicle complex, as well as into the oral lamellae. Mesocolomic ducts are paired on the dorsum and possess a thicker, more ciliated epithelium on ventral surface of the duct (Hyman 1959; Benito & Pardos 1997; Dilly et al. 1986c). Dilly et al. speculate that these ducts allow the expulsion and intake of water when the feeding apparatus is contracted and expanded. The posterior of mesocoelomic cavities abut another septum formed by contact with metacoelomic sacs. The metacoels are similar to mesocoels in that they are paired and separated by dorsal and ventral septa. These cavities lack openings to the exterior. Most of the coelomic space is occupied by digestive system or gonads, leaving little open space (Hyman 1959; Benito & Pardos 1997). These cavities partially extend into the posterior stolon.

8.2.4 Musculature Although pterobranch colonies are sessile, individual zooids and clusters can display a fair amount to of

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mobility. The arms and tentacles are active during feeding, the cephalic shield demonstrates considerable ability to move and contract, and the long stolon extending from the trunk can change its length several-fold through contraction and extension. There have not been comparative studies of musculature that compare colonies where a zooid is confined to a single tube (e.g., rhabdopleurids) to zooids that can freely move around the coenecium. Assessing how musculature varies with zooid independence would be interesting. As described in Hyman (1959), muscle layers are not well developed, but longitudinal muscle fibers are found just below the epidermis over much of the zooids body. Perhaps the most complex musculature in the pterobranch body is found in the cephalic shield. The longitudinal muscle tracks are well developed, except on the ventral side of the cephalic shield where muscles are not organized into distinct tracks. Nonetheless the cephalic shield still houses considerable myoepithelial tissue compared to other body compartments. Much of the muscular in the collar region is associated with the buccal cavity and arises from the septum between the collar and the trunk. Longitudinal muscles of the trunk, which appear to occur in two main bundles, originate near the basement membrane and can form a relatively thicker layer on the ventral side of the trunk. These tracks extend down into the elongated stolon. At the cellular level, muscle cells are myoepithelial in nature, with one end connected to the basal lamina, and mitochondria tend to be abundant within these cells. Of particular note, Ridewood (1907b) reported cross-striated fibers in pterobranchs. Hyman (1959) dismissed this report in her typical style, but Dilly et al. (1986c) confirmed cross striations in Cephalodiscus muscle. Myoepithelial cells use “zonulae adhaerentes” junctions for cell-to-cell connections.

8.2.5 Nervous system and sensory structures Pterobranchs, like enteropneusts, possess a rather diffuse and poorly centralized nervous system. In addition, their nervous system is mainly intraepidermal in nature, a condition typical of many smaller animals (Hyman 1959; Rehkämper et al. 1987). The most notable concentrations of nerves are in dorsal epithelium of the collar region near the base of the tentacular arms and in the tentaculate arms (Rehkämper et al. 1987; Stach et al. 2012). To a large degree, this placement makes biological sense as the feeding apparatus is the most active part of the zooid

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and food chemosensory abilities are needed to distinguish food particles from non-desirable particles. In agreement with Hyman, Rehkämper et al. report that nerve tracks that radiate to other parts of the animal originate from the collar ganglion and that these nerves show a distinct thickening on the ventral midline of the trunk. This dorsal nerve plexus also possesses a neuropile. Neural bundles in pterobranchs (especially in the dorsal collar nerve plexus) are not hollow (Hyman 1959), and given the intraepidermal nature of the nervous tissue, there is no structure that appears to be homologous to a chordate dorsal hollow nerve chord. Although little work has been conducted on the pterobranch nervous system, the general conclusion has been that it is much less complex and more diffuse than nervous systems in enteropneusts or even the hemichordate sister group, Echinodermata. Specific sensory cells are not known, and pterobranchs do not have eyes. Stach et al. (2012) provide reconstructions of the pterobranch nervous system using immunohistochemistry and 3D microscopy.

8.2.6 Digestive system Pterobranchs are suspension-feeding animals that use their ciliated tentacles and arms to clear algal cells and other small food items from the water. To carry out their feeding, they typically perch at coenecial openings and inflate their feeding apparatus. In species with one zooid per coenecial tube, they often sit with their cephalic shield, cupping the edge of the coenecium (Lester 1985). In other species such as C. gracilis and C. hodgsoni, zooids will climb stalks made near the coenecial opening. Rhabdopleurids extend their pair of arms in a V-like pattern, but cephalodiscids typically deploy their tentacles to make a feeding sphere (Lester 1985). By generating a current with lateral cilia (approximately 15 µm in size) located on the tentacles, pterobranchs use a local reversal of ciliary beat to place food items on the oral surface of the tentacle (Halanych 1993). These particles are carried down the length of the tentacle to a larger ciliated food groove on the arm. From there, particles are transported to the mouth in the collar region and ingested. Tentacles can display flicking behavior to help manipulate food particles and reject undesirable particles prior to reaching the mouth, indicating selectivity in feeding. The digestive system is composed of a mouth, pharynx (including the buccal region and gill pore regions), a stomach, and a U-shaped intestine. Food enters the mouth, which is located at the anterior ventral side of the collar, is protected by the cephalic shield, and has oral lamellae

with circular muscle that act as lips. The esophagus connects the pharynx to a stomach region that is characterized by thicker epithelium that contains more glandular and columnar-shaped cells. The buccal and pharyngeal regions appear to largely consist of monociliated and mucous-producing cells (Hyman 1959; Benito & Pardos 1997). As in acorn worms, a buccal diverticulum extends anteriorly from the dorsal side of the buccal cavity. This structure, or stomal chord, is composed of two sheets of epithelium and a discontinuous lumen that projects anteriorly into the cephalic shield. In Cephalodiscus, one pair of gill pores is found at the posterior end of the pharynx, but Rhabdopleura lacks any external gill opening. The gill region lacks collagen and cartilaginous structural elements, but vacuolated cells in this region have been postulated to provide support (Dilly et al. 1986a). Interestingly, epithelial cells on either side of the gill pore show distinct morphology with densely ciliated osmiophilic cells on one side and the vacuolated, supportive cells on the other side. The presence of the gill pore is likely to handle excess water flow into the mouth due to filter feeding (Harmer 1887; Gilmour 1979). A short esophageal tube connects the pharynx to an enlarged stomach region that can occupy much of the zooids trunk region. The stomach has a caecum, and the epithelium is remarkably uniform, especially in Rhabdopluera, with tall columnar cells that clearly serve a secretory function. The posterior of the stomach funnels into the intestine, which has a pronounced change in cell type. There are few secretory cells in the intestine whose function is mostly resorptive with microvilli and cilia becoming progressively less common moving toward the end of the digestive track (Benito & Pardos 1997). The anus is placed anteriorly on the dorsum of the trunk. The best available descriptions of the pterobranch digestive system can be found in Dilly et al. (1986a) and Benito and Pardos (1997).

8.2.7 Gas exchange system Given that pterobranch zooids are on a millimeter size scale, they presumably obtain much of their oxygen through diffusion. The tentaculate arms offer enormous surface area especially in Cephalodiscus, and as mentioned below, the arms contain a blood vessel and a mesocoelom that can help facilitate oxygen absorption and transport as well as passive excretion of waste products (e.g., carbon dioxide and ammonium). The ciliated gill pores in Cephalodiscus attracted much attention from early workers leading to their placement with acorn worms (Willey 1899a, b). By contrast,

8.3 Reproduction and development 

Rhabdopleura lacks these openings but has pharyngeal pouches that do not open to the exterior environment. The pores in Cephalodiscus are rather simple in nature. The size and feeding mechanics of pterobranchs suggests that the evolutionary maintenance of gill pores may have more to do with feeding than gas exchange. Specifically, the use of ciliated mechanisms to move particles to the mouth also entrains a fair amount of water that must be expelled, presumably after the food particle has been secured in the pharynx.

8.2.8 Excretory and circulatory systems In general, the blood vascular system of pterobranchs is rather simple, likely due to their diminutive size and large surface area that aids diffusion. The blood vascular system is composed mainly of small sinuses, or lacunae, that lack definitive walls (Hyman 1959; Dilley et al. 1986b; Benito & Pardos 1997). There are slightly better developed dorsal and ventral vessels that run mainly through the collar and trunk regions. However, the feeding apparatus contains well-developed blood vessels located within the mesocoelom (Benito & Pardos 1997, fig. 61). Moreover, cells can be seen to moving within these vessels when the feeding apparatus is fully expanded (Halanych, personal observation). Similarly, Dilly et al. (1986b) reports the presence of a fine granular material in the blood assumed to be blood pigment proteins. The most complex excretory and circulatory organs are contained in the glomerulus-pericardial complex, which is also found in enteropneusts. The glomerulus contains vessels that interact with a dorsal blood vessel to help filter blood. Dilly et al. (1986b) demonstrated the presence of monociliated podocytes from glomerular vessel wall push into the protocoel in Cephalodiscus. Pericardial contractions are thought to generate the pressure needed for the filtration of blood relative to the coelomic fluid. This filtrate is then passed to narrow tubes that exit through the protocoel pores of the cephalic shield.

8.3 Reproduction and development As might be expected of colonial animals, pterobranchs take advantage of both sexual and asexual reproduction. Pterobranchs have been described as both simultaneous hermaphrodites and dioecious, but distinguishing between the reproductive status of the zooid versus the cluster or colony is important. Hermaphroditic zooids

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have been reported (Rhabdopleura, C. nigrescens, and C. hodgsoni), but individual zooids are more commonly male or female. Colonies can house both sexes (e.g., Cephalodiscus gilchristi Ridewood 1908) (Ridewood 1908; Hyman 1959). Some colonies have been reported to contain “neuter” zooids (Hyman 1959; Benito & Pardos 1997); however, such reports appear unsubstantiated. To the best of our knowledge, there has never been a detailed assessment to determine if zooids are truly non-reproductive (as in bryozoan feeding zooids, or hydroid gastrozooids) or just not sexually mature. Similarly, some of the earliest reports (e.g., John 1931) discussed sexual dimorphisms between male and female zooids (e.g., C. hodgsoni). However, this interpretation need verification as multiple developmental states are present throughout some colonies of Cephalodiscus. Moreover, unhealthy or senescent zooids can lose arms (Halanych, personal observation), which may impact observations about how many arms mature zooids possess. In the roughly 140 years pterobranchs have been known, there have only been about five papers (e.g., Gilchristi 1917; John 1932; Stebbing 1970; Lester 1988a, b; Dilly 2014) focused on their reproductive and developmental biology. Sato et al. (2008) noted some seasonality to the reproductive state of Rhabdopleura compacta, and larvae have been collected from wild caught animals. Nonetheless, there is not a reliable, and easy to collect, source of gravid pterobranchs or pterobranch embryos, and thus their reproductive biology has largely remained an enigma. The comments below are drawn mainly from these early reports as well as Hyman (1959) and Benito and Pardo (1997), which are in large part compilations of previous sources. Perhaps the best information is for two Bermudan taxa (Lester 1988a, b; Dilly 2014) (currently recognized as R. normani and C. gracilis; see species entries in Section 8.5, “Phylogeny”) and C. nigrescens (John 1932; Dilly 2014), but this has been augmented with our own observations.

8.3.1 Reproductive system When gravid, reproductive organs occupy much of the trunk region. Rhabdopleura is reported to have a single gonad, whereas Cephalodiscus species have paired gonads that lie on either side of the dorsal midline. In both cases, the gonoduct(s) exit the trunk in an anterior dorsal position not far from the anus. Previous reports state that zooids can only be sexed by microscopy (a statement more recently reiterated by Benito & Pardos 1997), but this is incorrect. When zooids are not gravid, microscopic

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Fig. 8.4: Gonads in pterobranchs. Male zooid of Cephalodiscus fumosus showing white gravid testis through body wall of trunk region. Scale bar, 1 mm.

sections may be required to determine sex. However, when gravid, the sex of the zooid is often clearly visible. In the case of females, large eggs are visible though the trunk epithelium. R. normani possesses one large dark brown/ greenish egg (200 µm; Lester 1988a), and Cephalodiscus has one or two usually bright orange eggs. Similarly, gravid testes are often the typical color of sperm, white to off white (Fig. 8.4). Lester (1988a) gives an account of the ultrastructure of the gonad in R. normani. The ovary contains multiple oocytes, but typically only one is mature, per ovary, at any given time. The eggs are yolky and large (up to a third the length of the trunk, Dilly 2014), but a significant vitelline or yolk gland has not been well characterized. Dilly (2014) recently reported that the ovarian duct is oval in shape and thus larger than previously recognized. Previous reports debated if the ova could pass through the duct, or if ova were released by a tear in the epithelium (see discussion in Dilly 2014). Based on personal observation (Halanych, personal observation), both scenarios appear viable. In C. gracilis, ova can tear out of the dorsum of the trunk epithelium causing a major wound. The damage zooid will then turn into a brown body (see below). Given the lack of muscle in the gonoduct, muscular contractions of the trunk and the metacoelom acting as a hydrostatic skeleton likely aid the ejection of the mature eggs (Dilly 2014). Spermatozoa are typically elongate in form, but the number of mitochondria per cell appears limited

suggesting that pterobranch sperm have limited swimming capabilities. Whether fertilization is external or internal is not known. Dilly (2104) found mature sperm in the coenecia of Cephalodiscus colonies, but if these penetrate the female zooids is unclear.

8.3.2 Development The most complete, yet still limited, observations of development and metamorphosis are those of Lester (1998b) for Rhabdopleura and Gilchristi (1917), John (1931), and Dilly (2014) for Cephalodiscus. Early development appears to be radial holoblastic cleavage (Gilchristi 1917; Lester 1988a). Gastrulation has not been adequately described, and the early larvae superficially resemble planulae. R. normani can hold several embryos per tube in addition to the one zooid. Lester (1988b) depicts embryos as being held below the zooid, but we have also observed them to reside above the zooid (closer to the coenecial aperture) when they are more mature. The fate of developing larvae in Cephalodiscus appears to be more species specific. Ciliated larvae can be freely floating within the common coenecia as is the case in C. gracilis (Halanych, personal observation), or they can be fixed to coenecia (Dilly 2014; C. densus Schiaparelli et al. 2004). (Note that Dilly 2014 incorrectly asserts that development lacks a motile larval stage. He presumably makes

8.3 Reproduction and development 

this statement as pterobranch larvae are often brooded. However, if larvae are free of the tube, they will swim, and they have been observed swimming in the larger coenecia of C. gracilis and C. hodgsoni; Halanych, personal observation). Coelom formation has been described as schizocoelous (Dilly 2014), and the digestive track is initially a straight tube in the larvae (Hyman 1959; Lester 1988b). The planula-like larva develops a ventral depression or vestibule. At the time of metamorphoses, this ventral depression everts and is reported to turn into the ventral surface of the cephalic shield (Harmer 1905; Lester 1988b), but the description of this process is vague and needs further study. Following morphogenesis of the cephalic shield, the arms develop gradually, and organogenesis continues simultaneously with body region differentiation. In the case of Cephalodicus, we have observed freeswimming larvae that are at the planula stage in the large coenecia of C. hodgsoni. Presumably, they can escape the coenecium. However, Rhabdopleura can retain larvae with clear arm buds in the coenecia. Once a larva settles, it forms a primary zooid, or ancestrula, not unlike a bryozoan. Upon settlement, this sexually reproduced individual quickly forms a coenecium and begins to produce budding individuals. In the case of R. normani, Lester (1988b) reported a planula-like larvae settling outside the parent coenecium and having fully elongate arms within 57 hours and the ancestrula cocoon open within 100 hours. After the original progenitor, a colony forms by asexual development. In Rhabdopleura, stolons grow, bifurcate, and then produce additional zooids. Presumably something similar occurs in cephalodiscids that have one zooid per coenecial tube. However, in the case of several cephalodiscid species, clusters of zooids are formed through asexual development from the pedal disc, which also serves as an organ to attach the cluster to the substrate. As cephalodiscids have many pairs of arms, zooids can feed before growth is complete as long as they have one functional pair of arms. The pedal disc of such zooids usually display active budding. Asexual budding seems to happen as a fairly continuous process. Although Dilly (2014) describes budding in Cephalodiscus to some degree, much of the below description is based on Halanych’s observations made by tracking colonies of C. gracilis in Bermuda over a 3-month period. New buds develop as outgrowths of the pedal disk, which at first appear as a broad protrusion or out pouching from the pedal disc. The outpouching soon takes on a more circular shape and most of this tissue will become the cephalic shield of the new zooid. By

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the time the cephalic shield is distinguishable, a small stolon connects the shield to the pedal disc. Shortly after this stage, buds of the first pair of arms appear on the posterior portion of the circular region on either side of the midline. Note that the arms in adults are invaded by the mesocoelom, and thus the back half of this new bud region is presumably becoming the collar. As the zooids develop, pairs of arms are added from a medial position to a progressively more distal and posterior position. In adults, the insertion points of arms form a U-shape. Before the last pairs of arm buds are added, the primary arms already have tentacles and appear to be able to feed given the active cilia and tentacle and arm behavior. While the arm pairs are being added, the anterior region of stolon progressively thickens. Further microscopy is needed to more accurately determine origins of tissues that form the asexual bud. For convenience, the developmental stage for asexual buds of C. gracilis has been numbered by the pairs of developing feeding arms. Stage 0 is from the first signs of a bud through the stage where a circular structure on a small stolon is visible. Stage 1 is when the first pair of arms form as buds, stage 2 represents the formation of the second pair of arm, etc. When all arms and tentacles have completed development, the zooid is considered mature. This naming convention can be easily applied to other Cephalodiscus species. Dilly (2014) also hypothesizes about another type of reproduction carried out by an asexually developed dispersing larval form called a “black comma body” (also called a “brown body”). The structure in question is a brown club-shaped structure attached to the pedal disk with a ciliated epidermis. Dilly incorrectly argues that this body is grown with a cluster as an asexual larval form. Harmer (1905) previously noted that brown bodies are senescent zooids, which is a correct interpretation. Based on Halanych’s Bermuda observations, zooids that were damaged or that had shed their arms darkened in color and became a brown body. Interestingly, a small number of zooids that had shed their egg by rupture of the trunk epithelium also regressed to brown bodies. Moreover, the brown bodies range in size from the size of a mature zooid’s trunk to small skinny projections from the pedal disk. Light microscopy sectioning of these bodies possessed the internal organs of the trunk regions (mainly digestive tract). Smaller brown bodies appear to have an internal structure that is less organized. Brown bodies are apparently clusters resorbing damaged zooid tissue to redistribute the energy. The observations that brown bodies of well-fed clusters decrease over time is consistent with Harmer’s speculation and contra Dilly’s hypothesis.

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8.4 Distribution and ecology Pterobranchs are mainly distributed in the Southern Hemisphere and are most abundant and diverse in the cold waters surrounding Antarctica. In this region, they tend to live on the continental shelf at depths ranging from about 150 to 700 m. (Note that the Antarctic continental shelf is depressed to about 400 m due to the weight of ice.) However, as a group, pterobranchs range from intertidal to approximately 1500 m in depth. Other localities include the southern reaches of the African and South American continental shelves as well as off New Zealand and Australia. A few species, (e.g., R. compacta Hincks 1880, R. normani, Rhabdopleura recondita Beli et al. 2018, Cephalodiscus atlanticus Bayer 1962, and C. gracilis) are found in the Northern hemisphere. Only Rhabdopleura species are found around Europe. The Bermuda populations of R. normani and C. gracilis are of particular note as these are the best-studied pterobranchs due to the relative ease of collection. In Bermuda, these species occur on the same substrate in water just below the intertidal zone. However, given that both of these species are encrusting forms, considerable effort is needed to find and then to harvest material for study. Although most pterobranch species prefer cold water, tropical forms are known (Harmer 1905; Bayer 1962). As suspension feeders, pterobranchs are constrained to certain ecological limitations. For example, they are typically found in areas of high water flow and where the water is relatively free from large amounts of particulate matter (oligotrophic shallow waters or below the photic zone). Interestingly, most of the species that live in less than 150 m of water also seem to prefer calcium carbonate substrate. More specifically, they prefer the protected side of a calcium carbonate substrate in a high flow area. R. compacta around the British Isles can be found on the concave side of bivalve shell rubble of hash. R. normani in the Norwegian sea and surrounding waters prefers to live within reefs of the deep-sea coral Lophelia (Milne Edwards & Haime 1849). The recently described R. recondita is found in the Adriatic and Ionian seas, associated with the calcium carbonate skeletons of cheilostome bryozoans (Beli et al. 2018). The Bermuda and Florida forms of R. normani and C. gracilis prefer the undersides of coral rubble. These species do use other substrates but these are the best places to search for the animals. Little is known about the behavior or ecological interactions of pterobranchs with other species. As a group, even slight vibration will make them quickly retreat from a feeding posture. Presumably they are chemically defended, given the bright orange coloration of the Cephalodiscus zooids and that they have no structural defenses.

Anecdotally, Halanych has offered individual zooids of both genera to fish in Bermuda. The zooids were taken into the mouth and then spit out without further interest. However, the hypothesis of chemical defense remains largely untested. Interestingly, the encrusting forms are often over grown with other animals (e.g., tunicates, sponges, annelids), suggesting there is not a deterrent in the coenecium. On the other hand, the large coenecium of deep-sea Antarctic pterobranchs are often almost entirely free of epibiotic growth.

8.5 Phylogeny Pterobranchia is the sister clade to Enteropneusta, the acorn worms, within Hemichordata. Together with echinoderms, hemichordates form Ambulacraria within Deuterostomia (Halanych 1995; Cannon et al. 2014). (Note that the pelagic Planctosphaeroidea is likely just an enteropneust larval form and not a major clade of hemichordates.) The crown group Pterobranchia is composed of just two genera, Rhabdopleura and Cephalodiscus. Atubaria (Sato 1936), described as a third extant group of pterobranchs, is most likely a member of Cephalodiscus taken outside the tube (see Komai 1949; Halanych 1996; Tassia et al. 2016) and is considered a nomen dubium. Importantly, the alliance between modern pterobranchs and fossil graptolites has long been a subject of speculation (Allman 1872; Schepotieff 1905; Kozłowski 1938), and over the last 50 years, increasing evidence has been gathered to support this hypothesis (e.g., Kozłowski 1966; Crowther & Rickards 1977; Armstrong et al. 1984; Urbanek & Dilly 2000; Maletz 2014). More specifically, Rhabdopleura has been considered a living graptolite (Mitchell et al. 2013). Explicit phylogenetic approaches have been used to explore relationships among graptolite fossils and how they relate to Rhabdopleura. We point readers interested in graptolite phylogeny to the recent work of Maletz and colleagues (e.g., Maletz et al. 2009; Maletz 2014, 2017). Cephalodiscus, which also includes extinct Eocene species (Cephalodiscus lutetianus Abrard et al. 1950), is considered the sister lineage to the Graptolithina, which includes modern rhabdopleurids and their graptolite relatives. Phylogenetic relationships of modern pterobranch fauna have not been adequately explored. Formal phylogenetic analyses to date have included Cephalodicus or Rhabdopleura species as representatives for studies focusing on deep lineages. Relationships within the genera are poorly understood. Early works split the species of Cephalodiscus into four subgenera based on coenecial

8.6 Diversity 

characters: Idiothecia (Ridewood 1907b), Cephalodiscus (Demiothecia) (Ridewood 1907b) (see John & Muirhead 1951; Markham 1971), Orthoecus (Andersson 1907), and Acoelothecia (John 1931). Structure of the coenecium can vary with environmental conditions, and how indicative these structures are of phylogenetic history is not clear. Limited 18S nuclear and 16S mitochondrial ribosomal subunit gene data from five taxa (Cannon et al. 2013) show that C. hodgsoni and Bermudan C. gracilis are each other’s closest relative and a sister lineage to a C. fumosus, C. nigrescens, and C. densus clade. Both C. hodgsoni and C. gracilis are in the Cephalodicus subgenus that is characterized by a branching coenicum in which zooids occupy communal spaces. The other three species are either within Idiothecia (C. nigrescens) or Orthoecus (C. fumosus and C. densus). These subgenera form coenecia that are unbranched, and zooids are confined to a single tube. For Idiothecia, the newest tubes form at the apices of the branches, but tubes are unbranched in Orthoceus with the tubes arising at the edges of the colony’s base (Markham 1971). The Beli et al. (2018) study based on 16S and 18S ribosomal subunit gene data included the most Rhabdopleura lineages to date. An unidentified Rhabdopleura from the Gulf of Mexico is clusters with, and is genetically similar to, R. normani from Bermuda. A specimen from deep Icelandic waters clusters with, but is clearly genetically different from, R. compacta. This unidentified Icelandic Rhabdopleura could possibly be R. normani whose type locality is in Norway, but samples are needed from the type locality to confirm such speculation. The most recently described pterobranch species, R. recondita from Italy, is genetically distinct from other species and forms the sister group to the clade consisting of the undescribed Icelandic specimens and R. compacta. Most molecular data on pterobranchs have been collected for phylogenetic studies and includes transcriptomic data (Cannon et al. 2014; Simakov et al. 2015), ribosomal subunit gene data (Halanych 1995; Cameron et al. 2000; Cannon et al. 2009, 2013; Worsaae et al. 2012; Beli et al. 2018), and whole mitochondrial genome data (Perseke et al. 2011; Li et al. in press). Interestingly, some pterobranchs show a strong nucleotide bias in their mitochondrial genomes.

8.6 Diversity The diversity of pterobranchs is limited. Cephalodiscus contained 18 recognized extant species and 5 Rhabdopleura. Many of the species with large conspicuous

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coenecia live in cold waters (i.e., deep or high latitude). By contrast, many of the tropical forms are encrusting and can be very difficult to spot without a preconceived search pattern in mind. In addition, most of the formative work on pterobranchs occurred late in the 1800s or early 1900s when species descriptions tended to be less precise and informative compared to modern standards. As a result, some species (i.e., R. normani) have broad and unrealistic distributions given the environmental conditions. Thus, pterobranch diversity is certainly higher than currently recognized. Much of the taxonomic discussion concerning pterobranchs has focused on structures of the coenecia that tend to be used in diagnostic keys (e.g., Ridewood 1907a, 1920; Markham 1971) to distinguish between species. Although coenecial characters, and numbers of tentaculate arms, do permit some species discrimination in Cephalodiscus, as a rule coenecial morphology tends to be highly variable. As with other suspension or filter feeders living in high flow areas, the shape of the organism (in this case coenecia) can be highly influenced by water currents as the structure grows. In Rhabdopleura, younger tubes tend to be more translucent and have less pronounced annulations. Moreover, the degree to which the tubes are elevated from the substrate may be a function of water current strength. All pterobranch zooids tend to have the same general morphology, but detailed comparative work on the morphological diversity of the zooids, or development, is wanting. Alignment of such characters align with species boundaries is unknown, and perhaps, there are numerous cryptic species. Although pterobranch taxonomy and diversity needs to be addressed with a systematic approach, the difficulty of obtaining specimens and the poor quality of previously collected samples present a substantial challenge. Most taxonomic descriptions of pterobranchs are from the last half of the 1800s or the first part of the 1900s. Many earlier works dealt with describing general pterobranch morphology as they were a new and unknown lineage at the time. Thus, the taxonomic descriptions are, by today’s standards, vague and non-descript. Genus Rhabdopleura Allman 1869b. Discovered in Norwegian waters in 1866 by G. O Sars. M. Sars applied the name Halilophus mirabilis to the animal in 1869 in a faunal list but failed to provide a description making the name a nomen nudum. Allman’s (1869) name has been used since then. Zooids in Rhabdopleura are 1–5 mm in length and form pale brown to dark brown to blackish coenecia that only hold one zooid per erect tube. Zooids possess one pair of tentaculate feeding arms and gill pores are absent.

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Rhabdopleura annulata Norman 1921. The type locality is Three Kings Island, north of New Zealand in 183–549 m. Annulation on the tubes is more pronounced than in R. normani, but coenecial structure and organization is similar. Coenecium is pale brown in color. Rhabdopleura compacta Hincks 1880. Compared to R. normani, which typically forms creeping coenecia, this species builds its tubes close together to form a “compact crust” with several tubes placed side by side. This species was originally found on shells from “deep water” (recent collections were in 20 m depth; Satoh et al. 2008) off the coast of Antrim in the United Kingdom. Recent collections have been just south of Plymouth, UK. Rhabdopleura normani Allman 1869b. Type species. As the first pterobranch species to be formally described, information that diagnoses this species as unique from other pterobranchs is limited. However, this species usually forms a creeping coenecium, and the individual tubes are clearly annulated. New coenecia can be pale or colorless, but older coenecia are dark brown to a translucent black. The type locality is Lofoten Islands, Norway. The species’ distribution is likely the colder waters of the north Atlantic. The R. normani reported from Bermuda is morphologically very similar to individuals from in Norway. However, the likelihood of these being the same species is low. The Norwegian form occurs at depths roughly 30 to about 300 m in cold water, whereas the Bermudan form occurs at depths as shallow as 1 m in warm oligotrophic water. Moreover, the Bermuda form is genetically similar to zooids found in the Gulf of Mexico (Cannon et al. 2013). Direct morphological and molecular comparisons between these zooids from these localities are needed. Rhabdopleura recondita Beli, Cameron & Piraino 2018. This is a recently described pterobranch found in cheilostome bryozoan skeletons off the coast of Italy in 2–70 m. The zooids are slightly longer than those of R. compacta. The distal tips of the arms lack tentacles for approximately one third of their length and are darkly pigmented. The colony forms a creeping coenecium that grows within the bryozoan fuselli. The coenecium is reasonably well annulated, and molecular ribosomal data confirmed its uniqueness compared to other Rhabdopleurids. Rhabdopleura striata Schepotieff 1909. This species was found near the small town of Kankesanturai on the Palk Strait coast, in Sri Lanka (formerly Ceylon), in 2–3 m of water. In the description, Schepotieff repeatedly refers to how similar R. striata is to R. normani. He does note the tubes are more of a pale yellowish color rather than the darker color of R. normani.

Genus Cephalodiscus M’Intosh 1882. Discovered on the Challenger expedition in 1872 in the Straits of Magellan at about 448 m depth. Cephalodiscus has multiple tentaculate arms and forms coenecia that tends to be pale yellow to orangish in color. The coenecia are not annulated as is the case for Rhabdopleura. The zooids tend to be a dark brown to red to bright orange in color and can be 3–14 mm in size (excluding the stolon). The zooids have a stolon that connects to a pedal disk that serves as a zone of proliferation of asexually produced zooids. Within Cephalodiscus, the coenecium can take many forms from agglutinated structures impregnated with sand and mud to erect, translucent massive colonies. Markham (1971) provided a revised diagnosis for the genus. Cephalodiscus agglutinans Harmer & Ridewood 1913. As the name implies, this species builds tubes that are agglutinated with sand, sediment, and particles from other organisms (shells, urchin spines, etc.). The coenecium can be large, 10 cm, and the internal spaces are connected but retain a tubelike network. Whether Harmer and Ridewood, who originally described this species, thought it has Cephalodiscus or Idiothecia construction is unclear (although Markham 1972 called it Idiothecia). The type locality is Burdwood Bank south of the Falkland Islands. Zooids are on the larger side (approximately 7 mm) and typically have nine pairs of feeding arms but variation has been reported. Cephalodiscus atlanticus  Bayer 1962. Collected near Cay Sal Bank in the Straits of Florida in 1955 at 275 m depth, this species has coenecium construction of Cephalodiscus subgenus type and was noted for being similar to C. sibogae and C. gracilis (Bayer 1962). Projections and spines rise up for the common coenecial chambers. Recovered colonies are small in size (8000 m in depth. Tassia et al. (2016) reviewed the global distribution of recognized species,

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although the authors caution that hemichordate biodiversity remains under-described, so any such report will be an underestimate. Enteropneusts are benthic, living in burrows, under stones, among algal holdfasts, or on the surface of deep sea sediments (Hyman 1959; Osborn et al. 2011). Saccoglossus and Balanoglossus are particularly known for their burrowing behavior. Some enteropneusts ingest sediment, extract organic material during digestion, and then produce a characteristic fecal casting on the benthic surface. Not all enteropneusts produce these castings, however. There have been few studies making use of time lapse imaging to study the fecal castings of deep sea torquaratorid enteropneusts (Smith et al. 2005; Anderson et al. 2011; Jones et al. 2013). Many enteropneusts give off a characteristic odor, presumably from the bromo-organics they produce, which have been hypothesized to be unpalatable defensive compounds (Thomas 1972; Fielman & Targett 1995; King et al. 1995; ­Kicklighter et al. 2003). Kicklighter et al. (2004) found that S. kowalevskii was unpalatable to two sympatric species of fish because of its high concentrations of 2,3,4-tribromopyrrole, although the worm was still readily consumed by a sympatric crab. There are few studies on acorn worms’ species interactions. Some species of enteropneusts have been reported to live inside the burrows of other species, for example, G. ruficollis is found inside Balanoglossus carnosus burrows (Willey 1899c; Okuda 1939; Hyman 1959). Protozoan parasites, particularly gregarines, have been reported in the hepatic sacs of B. clavigerus, Harrimania kupfferi (Hyman 1959), and Glossobalanus minutus (Wakeman et al. 2014). A parasitic copepod in P. flava was described by Tung et al. (2014). Several species of ptychoderid have been observed to be bioluminescent, although the in vivo mechanism and the function of this response remain unknown (reviewed in Oba et al. 2017). Although ecological studies on adult enteropneusts are scarce, research on larval ecology is even more rare. Little is known about larval feeding or predation in tornaria larvae. One recent study found that growth rates in the tornaria larvae of Schizocardium sp. from Texas were adversely affected by exposure to chemically dispersed crude oil (Almeda et al. 2014). Ecological interactions of both enteropneust adults and larvae will benefit greatly from increased scientific scrutiny in the future.

9.5 Phylogeny Since the advent of molecular phylogenetics, there has been a consensus on relationships between the three

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major deuterostome phyla, Chordata, Echinodermata, and Hemichordata (see discussion in Halanych 2004; Swalla & Smith 2008; Kocot et al. 2010). Phylogenomics has further upheld Ambulacraria (Bourlat et al. 2006; Dunn et al. 2008; Lartillot & Philippe 2008; Philippe et al. 2009; Cannon et al. 2014; Simakov et al. 2015), and there is additional support for this clade from morphology (Cameron 2005; Ruppert 2005), and a combined analysis of ribosomal, mitochondrial, and nuclear protein-coding genes (Bourlat et al. 2008). The Ambulacraria hypothesis suggests that shared morphological features between hemichordates and chordates, such as pharyngeal gill slits and the postanal tail, were likely to have been present in the deuterostome ancestor. Given general agreement on the position of hemichordates within deuterostomes, focus has turned to resolving the relationships within Hemichordata. Historically, there were few explicit phylogenetic hypotheses, although traditional wisdom assumed pterobranchs and enteropneusts were reciprocally monophyletic taxa based on their disparate morphology and life history (Dawydoff 1948; Hyman 1959). Beginning with Halanych (1995), studies using 18S rDNA sequence data have recovered Enteropneusta as paraphyletic, with pterobranchs originating within the acorn worm lineage sister to Harrimaniidae (Cameron et  al. 2000; Bourlat et al. 2003; Cannon et al. 2009; Worsaae et al. 2012). In their 28S rDNA analyses, however, Winchell et al. (2002) found pterobranchs as sister to a monophyletic Enteropneusta, a result that has also been recovered by Cameron’s (2005) morphological cladistic analysis, and phylogenomics (Cannon et al. 2014). Within Enteropneusta, phylogenetic hypotheses have been lacking, although Hyman (1959) noted that harrimaniids were the least complex of the enteropneusts, followed by spengelids and ptychoderids. Aside from this type of subjective assessment, the earliest objective studies addressing hemichordate relationships included Halanych (1995), with 18S rDNA data from three hemichordate taxa, and Winchell et al. (2002), who included six hemichordates in a broader study on deuterostome relationships using 18S and 28S rDNA data. Cameron (2005) conducted a morphological cladistic analysis including all known hemichordate genera at the time, although resolution within Enteropneusta was poor. Cannon et al. (2009) specifically addressed hemichordate relationships using 18S rDNA and two mitochondrial markers; several subsequent studies have built upon the Cannon et al. (2009) data set (Osborn et al. 2011; Worsaae et al. 2012; Cannon et al. 2013; Cedhagen & Hansson 2013; Holland et al. 2013). Cannon et al. (2014) included 14 hemichordate species from all recognized families in a

phylogenomic analysis of data sets consisting of 162–299 genes. Generally, these studies have recovered traditional monophyletic enteropneust families with the exception of placing the unusual harrimaniid S. canadensis as sister to all other enteropneusts (Cannon et al. 2014; Li et al. under review). Saxipendium, which had been considered to ­constitute its own family, was also found to be nested within Harrimaniidae (Cannon et al. 2009; 2014; Deland et  al. 2010). In addition, ribosomal, phylogenomic, and mitochondrial data sets have suggested that Torquaratoridae is within Ptychoderidae (Cannon et al. 2013; 2014; Li et al. under review). Although represented by no more than two species in any molecular phylogenetic analysis to date, Spengelidae has been consistently recovered as sister to Torquaratoridae + Ptychoderidae. Harrimaniidae is sister to the remaining enteropneust families. Molecular phylogenetic analyses published to date have not contained sufficient species coverage to thoroughly test monophyly of most genera, nor the relationships of genera within families. A phylogenetic hypothesis for the relationships of described extant hemichordate genera is given in Fig. 9.14. In addition to the described species of enteropneusts, there are several undescribed taxa that have been characterized molecularly or documented photographically (Cannon et al. 2009; Anderson et al. 2011; Osborn et al. 2011; Cannon et al. 2013). Cannon et al. (2009) included sequence data from a tornaria larva that is not phylogenetically close to any other species, falling sister to the clade of Torquaratoridae + Ptychoderidae, but not within Spengelidae (Cannon et al. 2009, 2013; Osborn et al. 2013). In addition, there are at least six undescribed species of Torquaratoridae with published molecular sequence data (Cannon et al. 2009, 2013; Osborn et al. 2011), and many that have been photographed but not collected (Holland et al. 2005; Smith et al. 2005; Anderson et al. 2011; Osborn et al. 2011). Cannon et al. (2013) used molecular data to characterize a novel group of cold-water harrimaniids that is split into four clades, as well as a distinct Icelandic harrimaniid that does not phylogenetically align with any other genus with available sequence data.

9.6 Diversity In 2010, Deland et al. presented a revised taxonomy of Harrimaniidae based on historical collections, started over 100 years prior, from Ritter, Bullock, and Rao. These collections, composed of complete specimens, sections, notes, and micrographs, had been passed from Bullock to Burdon-Jones, who deposited the material at the

9.6 Diversity 

Cephalodiscus Rhabdopleura Stereobalanus Meioglossus Horstia* Protoglossus Harrimania Saccoglossus Mesoglossus Ritteria Saxipendium Schizocardium Spengelia* Willeyia* Glandiceps Balanoglossus Ptychodera Glossobalanus Yoda Torquarator* Tergivelum Coleodesmium Allapasus

 317

Pterobranchia

Harrimaniidae

Ptychoderidae

Spengelidae

Torquaratoridae

Smithsonian Institution upon his retirement. Bullock contacted Cameron to complete the work, and after Bullock’s death in 2005, several papers have now been published posthumously based on this material (Cameron et al. 2010; Deland et al. 2010; Cameron & Perez 2012; Cameron & Ostiguy 2013), substantially updating (and consolidating) enteropneust taxonomy and improving representation of morphological diversity within select acorn worm groups. At the time of this publication, the 108 described enteropneust species are classified in four families, with no ordinal level classifications. Diversity within the group is underestimated, with some authors estimating species numbers closer to 500–1000 (Appeltans et al. 2012). The four currently recognized enteropneust families are Ptychoderidae Spengel 1893, Spengelidae Willey 1899c, Harrimaniidae Spengel 1901, and Torquaratoridae

Fig. 9.14: Phylogenetic hypothesis for the described genera of Enteropneusta. Tree summarizes molecular phylogenetic results from Cannon et al. (2009), Osborn et al. (2011), Worsaae et al. (2012), Cannon et al. (2013), Osborn et al. (2013), and Cannon et al. (2014), with the addition of morphological data from Holland et al. (2005) and Deland et al. (2010). Genus names followed by an asterisk are represented by morphological data only.

Holland et al. 2005. In addition to these four families, Planctosphaeroidea is a rarely collected monotypic group known only as large, modified tornaria larvae of Planctosphaera pelagica. Although Planctosphaeroidea was once considered a separate class of hemichordate (van der Horst 1936; Hyman 1959), it is now commonly considered to be the larva of an unknown enteropneust (Hadfield & Young 1983). These planktonic organisms can measure up to 25 mm. They have been collected in the Atlantic and the Pacific, from depths of 75 to ~1000 m, but only about 30 individuals have been collected since their discovery in 1910 (Hart et al. 1994). Because of their uncertain affinities and taxonomic status, we have not included them in the classifications herein. In the following section, we briefly describe the four recognized families in the order they were described, and the genera that comprise them, also in order of year of

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description. This discussion is not intended to be a complete list of synonyms and collection sites. For further details, see Supplementary Table 1 from Tassia et  al. (2016). Many significant identifying characteristics of enteropneust species are internal morphology that must be observed via histological sectioning. In the following section, external anatomy has been emphasized, with the intention that this discussion may prove useful for nonspecialists to identify acorn worms in the field. References to more thorough internal anatomical descriptions are given under each genus. For a taxonomic key to the families of Enteropneusta, see the study of Deland et al. (2010), which also contains a key to the genera of Harrimaniidae. Cameron and Perez (2012) provided a key to the genera of Spengelidae; Cameron and Ostiguy (2013) provided a key to the genera of Ptychoderidae. Information for family and genus descriptions is from these sources unless otherwise indicated. Ptychoderidae Spengel 1893 Ptychoderidae includes 44 species in three recognized genera, Ptychodera, Glossobalanus, and Balanoglossus. Ptychoderids have been described as the most derived enteropneusts because of structures such as synapticles in the gill slits, pronounced regionalization of the trunk, and lateral septa (Hyman 1959). Ptychoderidae is defined by the absence of abdominal pores and the presence of lateral septa in the trunk. The primary and the secondary gill bars are connected via synapticles. The pericardium is simple, and there are dorsal nerve roots in the collar. Hepatic caeca are usually present, and the horns of the proboscis skeleton rarely extend past the anterior half of the collar (Cameron & Ostiguy 2013). Ptychoderids develop indirectly via tornaria larvae. P. flava has also been the subject of the majority of studies to date on indirect development among enteropneusts (e.g., Tagawa et al. 1998; Henry et al. 2001; Nakajima et al. 2004; Lin et al. 2016). More recently, B. misakiensis and Balanoglossus simodensis have been proposed as models for hemichordate indirect development, in part due to the ease with which these species can be reared in the lab (Urata & Yamaguchi 2004; Miyamoto & Saito 2007, 2010; Ikuta et al. 2009; Miyamoto et al. 2010). In molecular systematics studies, relationships between the three ptychoderid genera are extremely variable. Glossobalanus is non-monophyletic in several studies (Cannon et al. 2013; Osborn et al. 2013), suggesting some sequences may have been misidentified, or this genus requires systematic revision. P. flava, the first hemichordate to be described, is now one of two enteropneust species to have a complete published genome (Simakov et al. 2015).

Ptychodera Eschscholtz 1825 In Ptychodera species, the atrium opens via long slits that are exposed in their entirety to the outside. The genital wings are well developed. P. flava had the highest reported species distribution by Tassia et al. (2016), although the population and the species boundaries within Ptychodera are in need of further assessment. Urata (2015) analyzed 18S rDNA and mitochondrial 16S rDNA from P. flava specimens in Japan and found that these sequences were identical to sequences from P. flava collected in Moorea (Cannon et al. 2009), over 10,000 km away, suggesting that this species may indeed have an extensive geographic distribution. Balanoglossus Delle Chiaje 1829 As in Ptychodera, the genital wings in Balanoglossus are well developed. The primary distinction between Ptychodera and Balanoglossus in terms of external anatomy is that Balanoglossus has small pores opening to the atrium, rather than fully exposed long gill slits. Glossobalanus Spengel 1893 In Glossobalanus, there are genital ridges, rather than the genital wings found in other ptychoderid genera. The genital ridges usually extend into the hepatic region of the trunk. In most species, the hepatic sacs are consistent in shape and size and ordered in two rows. Gill slits are small in Glossobalanus. Spengelidae Willey 1899 Schizocardium, Willeyia, Glandiceps, and Spengelia comprise Spengelidae (20 species), which is defined by the presence of a digitlike projection, or “vermiform process”, at the anterior end of the stomochord. Spengelids additionally have a layer of circular muscle inside the layer of longitudinal muscle in the trunk, and the horns of the proboscis skeleton are long, generally extending through the entirety of the collar. They typically do not have dorsal nerve roots arising from the collar cord. Spengelids are described as having a combination of ptychoderid and harrimaniid features. For example, Spengelia and Schizocardium possess gill slit synapticles, whereas Glandiceps and Willeyia do not (Hyman 1959). In species where development is known, spengelids develop indirectly via tornaria larvae (e.g., Glandiceps—Rao 1953, Urata et al. 2014; S. californicum— Gonzalez et al. 2017). In general, this group contains many rare and poorly studied species; thus, there are few studies on spengelids. Illustrating this, a specimen of Glandiceps abyssicola, not seen since its discovery on the Challenger expedition in 1873, was recently rediscovered (Holland et al. 2013). However, there has been progress on develop-

9.6 Diversity 

mental biology in S. californicum (Gonzalez et al. 2017) and has gained attention as an emerging model species for indirect developing enteropneusts. Schizocardium Spengel 1893 The digit-like projection at the anterior end of the stomochord is very long in Schizocardium. The gill slits are also very long, extending almost the entire circumference of the pharyngeal trunk, leaving only a narrow hypobranchial strip of digestive pharynx. Synapticles are present, there are no dorsal gonads in the branchial region, and the hepatic sacs are pronounced. In addition, the pericardium bifurcates anteriorly into long tubes, and these are surrounded by a paired glomerulus. Glandiceps Spengel 1891 Synapticles, nerve roots, and hepatic sacs are absent. The ventral part of the pharynx is well developed. Glandiceps has an unusual swimming behavior, and swarms of swimming acorn worms have been described near Java (Spengel 1909) and in the Seto Inland Sea of Japan (Ikeda 1908; Yoshimatu & Nishikawa 1999). Urata et al. (2012) observed swimming behavior in cultured specimens of G.  hacksi and found that the worms could readily leave their burrows and swim by contorting their proboscis into a pear shape, flattening their trunk dorsoventrally, and wiggling their trunks. Worms swam on average 71 seconds, with the longest observed swimming time of 165 seconds (Urata et al. 2012). Spengelia Willey 1898 Synapticles are present, hepatic sacs are absent, and nerve roots may be present or absent. Pericardial diverticula are short, and there are dermal pits in the genital region. Willeyia Punnet 1903 The collar in Willeyia is longer than it is broad. Synapticles, nerve roots, and hepatic sacs are all absent. The pericardial diverticula are short, and dorsal gonads are absent. Harrimaniidae Spengel 1901 Harrimaniidae (40 species) is composed of members of the genera Saccoglossus, Harrimania, Stereobalanus, Protoglossus, Mesoglossus, Ritteria, Saxipendium, Horstia, Meioglossus, and Xenopleura. Harrimaniids are defined by the absence of many features; including circular muscles in the trunk, lateral septa, gill slit synapticles, and hepatic caeca (Deland et al. 2010). Harrimaniids have direct development, with a nondescript larval stage bearing little resemblance to tornaria. S. kowalevskii has become a well-known study organism

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for developmental work (Colwin & Colwin 1949, 1953, 1962; Stach & Kaul 2011; Kaul-Strehlow & Stach 2013) and developmental gene expression work (Lowe et al. 2003, 2006; Aronowicz & Lowe 2006; Lowe 2008; Darras et al. 2011; Green et al. 2013). These studies have yielded important insights by comparing hemichordate and chordate development. The S. kowalevskii genome is now available (Simakov et al. 2015). Saccoglossus Schimkewitsch 1892 Saccoglossus is most recognizable for the extremely long proboscis found in most of its members. Typically, at the proboscis, length is at least twice the width, but it may be much longer, and the proboscis has a middorsal longitudinal groove. The collar is generally very short relative to the proboscis. The longitudinal muscle fibers of the proboscis are arranged in concentric rings (Fig. 9.6 A). Harrimania Ritter 1900 Harrimania’s proboscis is cone shaped and is slightly longer than it is broad, the longitudinal muscles of the proboscis are arranged in radial plates (Fig. 9.6 B), and there are often two proboscis pores. The collar is broader than it is long, and the horns of the proboscis skeleton are very long, even reaching into the trunk in some species (Cameron 2002; Deland et al. 2010). The gonads form rows of simple sacs dorsally and laterally (Deland et al. 2010). Stereobalanus Spengel 1901 Stereobalanus is an unusual genus with four genital regions (two dorsolateral and two ventrolateral) immediately posterior to the collar. Broad gill openings with externally visible gill tongues are found between these paired gonad regions, opening to a single fused atrium (Reinhard 1942). The longitudinal musculature of the proboscis is arranged in radial plates (Fig. 9.6 B). Xenopleura Gilchrist 1925 This genus is described from a single specimen from the southern tip of Africa and is reportedly viviparous. Xenopleura is characterized by medullary folds in the dorsal trunk. Longitudinal muscles in the proboscis are diffuse (Fig. 9.6 C), and there is a single proboscis pore. Protoglossus van der Horst 1935 There is a deep dorsal groove at the posterior end of the short and conical proboscis in Protoglossus. In addition, the preoral ciliary organ is prominent and horseshoe shaped. The proboscis longitudinal musculature is radial (Fig. 9.6 B), and there is a single proboscis pore on the left side. The horns of the proboscis skeleton extend to the

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posterior of the collar, which creates ridges on each side of the buccal cavity. The anterior edge of the collar is ruffled (Deland et al. 2010). Saxipendium Woodwick & Sensenbaugh 1985 Commonly called “the spaghetti worm”, Saxipendium coronatum is a member of deep sea hydrothermal vent communities. The proboscis skeleton is crown shaped in cross section, with long horns that curve backward. The longitudinal muscles in the proboscis are diffuse (Fig. 9.6 C). Deland et al. (2010) formally synonymized Saxipendiidae (Woodwick & Sensenbaugh 1985) into Harrimaniidae after molecular phylogenetic analyses demonstrated that S. coronatum belonged in this group (Cannon et al. 2009). A second species, Saxipendium implicatum, was described in Holland et al. (2012) from seamounts in the deep sea off the central coast of California. Horstia Deland et al. 2010 In Horstia, the proboscis is quite short and rounded, with longitudinal musculature arranged in radial plates (Fig. 9.6 B). The gonads project from the trunk surface as modules. The gill pores are found on an elevated ridge, and the trunk itself narrows posteriorly from the collar (see Fig. 9.2 A, Deland et al. 2010). Mesoglossus Deland et al. 2010 Based primarily on Ritter’s unpublished manuscript written ca. 1900, Deland et al. (2010) erected the genus Mesoglossus to contain those species formerly of the genus Saccoglossus where the longitudinal musculature of the proboscis is arranged diffusely (Fig. 9.6 C), rather than in concentric rings (which is a defining character of Saccoglossus) or radial bundles. The proboscis is additionally approximately twice as long as it is wide, without a dorsal groove. Only lateral gonads are present, no dorsal gonads. There is one proboscis pore, generally on the left. Ritteria Deland et al. 2010 Ritteria has a short proboscis, and the longitudinal muscles of the proboscis are dispersed (Fig. 9.6 C). There is a single well-developed proboscis pore on the left side of the proboscis stalk, which is very reduced. The collar is wider than it is long. There are two pairs of dorsolateral genital ridges, and the branchial pores are found in a groove between these ridges. Meioglossus Worsaae et al. 2012 A genus of meiofaunal harrimaniid acorn worm, Meioglossus, has been described from material found in Bermuda

and Belize (Worsaae et al. 2012). The genus is characterized as microscopic, interstitial, and having a completely ciliated body. The proboscis is elongated and has a pair of proboscis pores at the base. The collar region is more than half the length of the proboscis. The sole described species, M. psammophilus, reaches a maximum body length of 0.6 mm, making it the smallest known enteropneust (Worsaae et al. 2012). Torquaratoridae Holland et al. 2005 Torquaratoridae was described in 2005 based on morphological analysis of a few specimens collected in the deep northeastern Pacific (Holland et al. 2005). This family now contains seven species in five genera, Torquarator, Tergivelum, Allapasus, Yoda, and Coleodesmium. To date, all torquaratorids have been found at 350 m or greater. Broad-collared acorn worms have been photographed in the deep sea since the 1960s (Bourne & Heezen 1965; Ewing & Davis 1967), but due to the extreme fragility of these animals, collecting intact specimens for species description has not been possible until recent advances in deep sea remote-operated vehicles (ROVs) (Osborn et al. 2011). Molecular phylogenetic results have indicated that Torquaratoridae is sister to, or within, Ptychoderidae (Cannon et al. 2009, 2013, 2014; Holland et al. 2009; Osborn et al. 2011; Worsaae et al. 2012). Torquaratorid life history is still poorly understood, although some species demonstrate unusual characteristics relative to their shallower cohorts. For example, videos taken by ROV have shown A. aurantiacus both burrowing and drifting above the benthos, indicating a possible benthopelagic life history (Holland et al. 2012). Torquaratorids are characterized by a broad proboscis and collar, and the proboscis skeleton is either absent or greatly reduced to a small plate. Synapticles are absent, and there are prominent hepatic caeca. The adult stomochord is separated from the buccal cavity of the collar or is otherwise absent (Holland et al. 2005; Osborn et al. 2011). Torquarator Holland et al. 2005 As the first described genus of Torquaratoridae, many of the characters used in the diagnosis of Torquarator bullocki, such as the relative breath of the collar, the absence of synapticles, etc., have subsequently been shown to be diagnostic of the whole family. Uniquely, T. bullocki is described as having a proboscis skeleton with very short anterior and posterior horns, which makes this the sole example of Torquaratoridae where the proboscis skeleton is at all elaborated beyond a simple plate. Unlike other described species in this family, there is no molecular sequence data for T. bullocki.

Literature 

Tergivelum Holland et al. 2009 There are currently two described species of Tergivelum, T. baldwinae and T. cinnabarinum. The primary external diagnostic character of Tergivelum is paired “black veils” that run along 30%–50% of the length of the trunk immediately following the collar. These structures were lost in collected specimens of T. cinnabarinum but were evident in images taken prior to collection. Internally, this genus is characterized by prominent buccal muscles on the left and right side of the mouth, which are strikingly robust in contrast to the remaining musculature. The two species are morphologically very similar and are distinguished by color, with T. baldwinae being dark brown and T. cinnabarinum named for its red, cinnabar coloration. Allapasus Holland et al. 2012 The proboscis complex in this genus is located in the proboscis stalk, more posteriorly than in other enteropneust families, and contains a plate-like proboscis skeleton without skeletal horns and no pericardial sac. In the female holotype specimen of A. aurantiacus, numerous ovaries are located in lateral wings that run the length of the trunk. Each ovary holds a single primary oocyte and is contained in a pouch of epidermal tissue that is connected to the lateral wing tissue by a thin stalk. A. aurantiacus and A. isidis are very similar, with primary differences in the descriptions relating to the fact that the holotype of A. aurantiacus is female whereas the A. isidis holotype is male. The two species differ in color, however, with A. aurantiacus being orange and A. isidis yellow. There are no observations of either of the two species of Allapasus leaving a fecal trail. Behavioral observations are limited to A. aurantiacus, which has been observed partially burrowed in sea floor sediments. This burrowing may be facilitated by the musculature in the proboscis and collar, which are both more robust than in other torquaratorids. Prior to collection, the A. aurantiacus holotype was recorded drifting a few centimeters above the sediment, apparently after voiding its gut contents, with no evidence of muscular undulations. Yoda Priede et al. 2012 The name Yoda comes from the shape of elongated lateral lips that account for two thirds of the width of the collar, and taper to points, resembling the ears of the Star Wars character, Yoda. The sole described species of this genus, Y. purpurata, is dark reddish purple in color. Specimens have been observed fully exposed on the seafloor surface, creating an irregularly meandering fecal trail.

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Coleodesmium Osborn et al. 2013 The sole described species of Coleodesmium, Coleodesmium karaensis, has a unique tubular sheath in the proboscis skeleton surrounding the collar nerve cord. In contrast to Allapasus, which is otherwise similar in general outward shape, the musculature in the proboscis and collar is weakly developed. The single collected specimen is a brooding female with embryos at differing stages of early development located on the surface of the pharyngeal region. This species was collected in the Russian Arctic at a depth of about 350 m, marking the shallowest reported collection of a torquaratorid enteropneust.

Literature Almeda, R., Bona, S., Foster, C.R. & Buskey, E.J. (2014): Dispersant Corexit 9500A and chemically dispersed crude oil decreases the growth rates of meroplanktonic barnacle nauplii (Amphibalanus improvisus) and tornaria larvae (Schizocardium sp.). Mar. Environ. Res. 99: 212–217. Anderson, T.J., Przeslawski, R. & Tran, M. (2011): Distribution, abundance and trail characteristics of acorn worms at Australian continental margins. Deep-Sea Research Part II Top. Stud. Oceanogr. 58(7–8): 970–978. Annona, G., Holland, N.D. & D’Aniello, S. (2015): Evolution of the notochord. EvoDevo, 6(1): 30. Appeltans, W., Ahyong, S.T., Anderson, G., et al. (2012): The magnitude of global marine species diversity. Curr. Biol. 22(23): 2189–2202. Aronowicz, J. & Lowe, C.J. (2006): Hox gene expression in the hemichordate Saccoglossus kowalevskii and the evolution of deuterostome nervous systems. Integr. Comp. Biol. 46(6): 890–901. Balser, E.J. & Ruppert, E.E. (1990): Structure, ultrastructure, and function of the preoral heart-kidney in Saccoglossus kowalevskii (Hemichordata, Etneropneusta) including new data on the stomochord. Acta Zool. 71(4): 235–249. Barrington, E.J.W. (1940): Observations on feeding and digestion in Glossobalanus minutus. Q. J. Microsc. Sci. 82: 227–260. Barrington, E.J.W. (1965): The Biology of Hemichordata and Protochordata. W.H. Freeman, San Francisco. Bateson, W. (1885): The later stages in the development of Balanoglossus kowalevskyi, with a suggestion as to the affinities of the Enteropneusta. Q. J. Microsc. Sci. 25: 81–122. Bateson, W. (1886): The early stages in the development of Balanoglossus. Studies from the Morphological Laboratory in the University of Cambridge 2: 131–160. Benito, J. & Pardos, F. (1997): Hemichordata. In: Harrison, F.W. & Ruppert, E.E. (eds.) Microscopic Anatomy of Invertebrates, pp. 15–101. Wiley-Liss, New York. Bourlat, S., Nielsen, C., Lockyer, A., Littlewood, D. & Telford, M. (2003): Xenoturbella is a deuterostome that eats molluscs. Nature 424(6951): 925–928.

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Index Acanthocephala 55, 59, 61, 83 acetylcholine 231, 241 acid hydrolysis 243 Acoelothecia 293, 295 actin 238 adhesive organs 240 adhesive papillae 208 adhesive structures 178 Aeropsis 151 agamete 4 Agassiz-stage tornaria 313 Aglaophamus 154 Aidanosagitta 171, 176, 178, 185, 186, 189, 190, 193, 237 alanine 249 Allapasus 299, 300, 306, 320, 321 alveolar tissue 199 Ambulacraria 292, 299, 316 Amiskwia 165, 173 amoebocytes 312 amoeboid cells 132, 133 Amphilepis 151 Amphipholis 36 Amphiura 36 Ankalodus 165 annex secretory gland 251 apical organ 146 archenteron 266, 267, 314 Archeterokrohnia 165, 177, 180, 186, 191, 237, 240, 249 Artemia 45 Arthropodaria 152 Ascidia 36 Ascopodaria 152 asexual reproduction 46 Astarte 35 Atubaria 292 axial cell 3, 4, 5 axoblast 4 Balanoglossus 299, 302, 307, 313–315, 318 Barentsia 111–113, 116–123, 125, 130–138, 140, 143, 148, 150, 152, 153, 156 baroreceptor 213 Bathybelos 177, 179, 183 Bathyspadella 165, 176, 177, 179, 180, 191, 194, 205, 249 Bdelloidea 55, 59, 83 binary fission 46 biological rhythms 194 bioluminescence 165 biomass 163 birefringent granule 43 blastopore 170, 261, 267 blood pigment 312 blood pigment proteins 289 Brachionus 267 brachyury 170, 256, 264 https://doi.org/10.1515/9783110489279-010

brood chamber 47, 90, 92, 102, 104, 140, 141 brooding 7, 314, 321 brood pouch 264 Bryozoa 107, 108, 111, 149 buccal diverticulum 288 buccal funnel 88, 90, 91, 95, 97, 99, 101, 126, 127, 137 budding 47, 155 buds 112, 113, 135, 136, 148, 154, 155, 291 buoyancy 164, 203, 242, 245 Burgess Shale 148, 165 Caecosagitta 165, 183, 191 Calispadella 177, 181, 192, 193 calotte 2, 3 Cambrian 165 cannibalism 259 Capinatator 165 capitate swellings 129 carbon flux 164 carotenoid pigments 165 castration 37 cave-dwelling 165 cell types 43 cephalic shield 284, 286–288, 291 chemoreception 220, 309 chemoreceptor 225 Chengjiang biota 148, 165 Chitaspis 152 chitin 205 Chlorella 45 chondroid tissue 305 chordoid larva 90, 91, 94, 97, 99, 100, 102, 104 chordoid organ 91, 94, 97, 98, 100 chordoid tissue 199 ciliary beating 20 ciliary fence organ 167, 168, 203, 216, 234 ciliary girdle 141 ciliary gliding 45 ciliary locomotion 21 ciliary slit receptors 216 ciliary tuft organ 167, 216, 219, 225, 263 Ciliocincta 11, 12, 18, 35 cleavage 3, 21, 35, 48, 82, 107, 141, 150, 249, 261, 266, 290, 314 cloaca 70, 77, 81 coelom 170, 245, 247–249, 251, 253, 255, 264, 266 coelom formation 170 coelomocytes 304 coelomoducts 249, 251 coelomogenesis 249, 261, 264 coelothel 170, 242, 245, 265 coenecium 283, 284, 288, 291, 292, 294, 295 Coleodesmium 320, 321 collar 284, 286, 299, 300, 302, 307–309, 312, 313, 318 collar coelom 300, 305, 306, 308

328 

 Index

collar cord 307, 318 collarette 177, 178, 199 collar nerve cord 321 collar region 283 colonial 111–115, 119, 121, 122, 129, 132, 133, 135, 136, 152, 289 colony 155, 283, 291, 295 Columbella 36 commensal 87, 151 Conocyema 3, 6 continental slope 189 copulation 11, 21–23, 35, 82, 106, 259 Coriella 111–114, 118, 152, 153, 156 corona ciliata 164, 166, 168, 178, 182, 203, 212, 216, 218–220, 225, 227, 265 Cotyledion 148, 149 Cryogenian 165 cryptic species 108, 178, 179, 293 Cryptomonas 45 crystal cells 43 cuticle 14, 15, 19, 57, 59, 70, 88, 94, 95, 111, 114–116, 123, 126, 130, 144, 146, 149, 157, 167, 203 Cyclatella 152 cylinder cells 43 cytoskeleton 20 Daphnia 261 Decipisagitta 177, 185, 186, 188, 191, 196 Deltoideum 148 dense layer 59, 61–64 desmosomes 20 deuterostomy 173 Dicyema 2, 3, 6–8 Dicyemennea 3, 6 Dicyemodeca 3 diet 163 Dinomischus 148 DNA bar coding 178 Dodecadicyema 3 dorsal antenna 67, 69 dorsal glands 93, 94 downstream collecting 129 Drosophila 261 duplicated ribosomal cluster 170 dwarf male 97, 98, 100–106, 149 Ediacaran 165 embryogenesis 3, 4, 247 embryonic development 35, 82 embryophore 136, 137, 140 Emschermannia 111, 112, 152–154 Endemism 193 endocytosis 243, 245 enterocoely 263, 265, 304 Eognathacantha 165 epibiont 111, 112, 151, 154, 292 epicuticle 94, 95 epipharynx 72–74, 84 Eukrohnia 164, 165, 168, 173, 178, 182, 189, 191, 193, 196, 205, 214–216, 237, 240, 249, 261, 264

Eurotatoria 83 exocytosis 197, 201, 256 extracellular matrix 21, 131, 132, 196, 200, 227, 233, 247, 255, 260 eye 178, 212–216, 223, 225, 265 eye pigment 182 eye spots 146, 309, 315 fecal pellets 164 fecal trail 321 fecundity 194 feeding 31, 34, 70, 128, 145, 148, 152, 163, 242, 245, 288 feeding arms 294 feeding rate 163, 194 feeding stage 87, 91–93, 95, 97, 99, 103, 104, 107, 149 Feldmannia 45 Ferosagitta 163, 171, 176, 178, 185, 186, 189, 190, 193, 199, 205, 216, 223, 225, 231, 234, 237, 238, 256, 259, 260, 262 ferritin 243 fertilization 21, 35, 37, 47, 48, 81, 106, 140, 256, 259, 260, 263, 267, 290, 312, 314 fiber cells 43 filter-feeding 131 filtration barrier 133 Flaccisagitta 176, 185, 186, 188, 190, 191, 193, 196, 237 flask-shaped cells 150 FMRFamide-like neurotransmitters 99 food availability 194 food supply 188 food transport 128 foot gland 112 foot groove 112 frontal organ 146, 147, 155 fulcrum 72, 73, 74, 84 gastrotroch 143 gastrula 313 gastrulation 142, 170, 245, 249, 261, 264, 266, 290, 314 gene rearrangements 41 genetic lineages 50 genetic toolkit 51 genital pore 13, 21, 22, 23 genital wings 302 genome 51 genome duplication 179 germline 261 giant fibers 223 gill pores 283, 284, 286, 288, 320 gill slit 302, 305, 309–313 Glandiceps 303, 305, 318 glial cells 223 glomerulus 299, 300, 311 glomerulus-pericardial complex 289 Glossobalanus 303, 307, 310–312, 315, 318 glycine 249 glycocalyx 114, 143, 144 glycogen 21 glycoproteins 242 Gnathostomulida 69 gonocoelom 249

Index 

gonocoelomoducts 249 Gonypodaria 152 graptolite 283, 292 grasping spines 166, 167, 178, 203, 205, 207, 211, 225, 231, 233, 259 gravity sensing 43 gregarines 315 growth zone 135 Haplozoon 1 Harrimania 306, 315, 319 hatchling 229, 265 heart-kidney complex 300 Heider-stage tornaria 313, 315 hemal system 170, 245, 247 Hemispadella 177, 181, 192, 237 hemocoel 114, 131 hepatic caeca 303, 318–320 hepatic saccules 302 Heteranomia 37 heterocoely 170, 249, 263 heterogony 37 Heterokrohnia 165, 173, 176, 177, 180, 185, 191, 205, 237, 240, 251 hibernacula 135, 136, 148, 156 Hoilungia 51 homeodomain sequences 172 hood 166, 207, 211, 225, 231 Horstia 306, 319, 320 Hox gene 170, 172, 256, 267 hybridization 259 hydroskeleton 75, 238 hydrostatic skeleton 290 Hyponeuria concept 251 hypopharynx 72, 73, 84 Idiothecia 293–295 immune reactions 304 incus 72 infection specificity 36 infusoriform 2, 3, 7 infusorigen 5, 8 insemination 37 internal budding 87, 88, 100, 102, 104 internal fertilization 37, 106 Intoshia 11–16, 18–27, 30–32, 34, 35, 37 intraepidermal layer 55 ion regulation 134 joint infection 37 Kaili Biota 148 keratin 199 Krohnitta 173, 183, 188–191, 193, 237 Krohnittella 177, 179, 182, 192, 193, 205 Krohn-stage tornaria 315 lacunar system 131 Lacunifera 150 lamina fibroreticularis 131 larva 11, 35, 37, 111, 137, 141, 142, 146–149, 154, 156, 290

 329

lateral antennae 69 lateral gland 93, 94 Lepeta 35, 36 Leptodora 213 Leptoplana 35 life cycle 11, 37, 38 ligamenta 73 lime-twig glands 118 Lineus 31, 35 lipophil cells 43, 45 lophophore 287 Loxocalyx 111, 152 Loxocorone 152–154 Loxokalypus 111–114, 152, 153, 155 Loxomespilon 152–154 Loxomitra 152–154 Loxosoma 112, 115, 121, 125, 127, 142, 143, 146, 147, 151–154 Loxosomatoides 111–114, 136, 152, 153, 156 Loxosomella 112, 115–118, 120, 121, 123–125, 127, 133, 135, 140, 141, 143, 145–148, 150–155 Loxosomina 153, 154 Loxostemma 152 Lucinoma 37 Macrorhynchus 30, 35, 36 magnesium inositol hexaphosphate 2 malleus 72–74, 84 mammary organ 140 manubria 72, 73 Maotianshan shale 165 marine food chain 165 marsupial sacs 261, 264 mastax 55, 61, 65, 68–70, 72–74 Mazon Creek 165 Medaka 261 meiofaunal 320 Meioglossus 299, 310, 319, 320 mesenteries 304 mesocoel 286, 287, 289, 304, 305 mesoderm 233, 245, 249, 256, 261, 263–267 Mesoglossus 306, 319, 320 Mesosagitta 171, 176, 177, 185, 190 mesosome 299 metacoel 287, 290, 304, 305 metagenesis 37 metaheterogony 37 metamorphosis 142, 143, 145, 147–149, 290, 315 metanephridial 249 metasome 299 metatroch 143 Metchnikoff-stage tornaria 313, 315 Microcyema 3, 6 Micrognathozoa 108 microsatellites 179 Midichloriacaea 43 mitochondrial genomes 51 mitogenomics 171 Monogononta 59, 61, 83 myoepithelial cells 233 myosin 238

330 

 Index

Myosoma 111, 114, 135, 152, 153, 156, 157 Mytilus 36 myxozoans 1 Natica 35, 36 Nebalia 55, 59, 70, 81, 82 neck gland 117 nematogen 1, 2, 4 Nematostella 261 Neosabellaria 35, 37 neotroch 313, 315 Nephrops 87, 88, 93 nephrostome 249, 253 Neresheimeria 1 neurocord 308 neuroendocrine vesicles 212 neurogenesis 256, 265 neuromuscular innervations 170 neurotroch 313, 315 neurulation 307 notochord 300, 310 nuchal organ 213, 220 Occulosagitta 176 ocelli 213 Oesia 165 oil vacuoles 196 ommatidia 215 Onoba 36, 37 oocyte 5, 15, 27, 47, 48, 58, 64, 77, 81, 90, 93, 100, 104, 106, 107, 136–140, 164, 248, 255, 261–264, 290 oogenesis 5, 80, 136 oogonia 80, 136–138, 248 oogonium 5, 8 Ophiotrix 36 Ophiura 36 opisthotroch 315 opsin genes 309 optical nerve 166 organogenesis 291 Orthoecus 293–295 osmoregulation 134 oxycline 195 oxygen minimum zone 188 Pandora larva 87, 90, 91, 97, 99, 101 Paramecium 20 paramyosin 238 Parasagitta 163, 167–169, 175–178, 184, 185, 190, 191, 193, 194, 196, 201, 205, 216, 227, 228, 237, 245, 247, 249, 253 Paraseison 55–57, 59, 62, 66–70, 72–76, 81–84 Paraspadella 179, 181, 192, 193, 208, 237, 238, 240, 263 Parhyale 261 parthenogenesis 37, 81 Paucijaculum 165 pedal disc 154, 285, 291 pedal glands 62, 69, 142, 147 Pedicellina 111, 116, 119, 121–123, 125, 134, 136, 137, 143, 147, 152, 153, 156, 157 Pedicellinopsis 111, 114, 152, 153, 156

Pedicellinosis 113 pericardium 1, 286, 300, 305, 307, 311, 314, 318 peritoneocytes 248, 253 peritoneum 304, 312 peroxidase 243 phagocytosis 45 Phakelodus 165 Philina 37 Phline 36 photoreception 309 photoreceptor 146, 213–215 phototaxis 143, 309 phragms 167, 178, 182, 183 pinocytosis 45, 312 placenta 140 Plagiostomum 152 planula larva 49 plasmodium 11, 18, 27, 29–33, 36–38 plasticity 178 Platynereis 261 plexus 201, 203, 221, 229, 233, 234, 240, 241, 288, 307, 312 plug cell 136 Podocoryna 49 podocytes 249, 289 Pododesmus 36 Poliometra 151 polyembryony 150 polymorphism 177 Polyzoa 108, 111, 149 population density 188 pore complex 118 predation 194 predators 48, 164, 231 prevalence 1, 2 prey 163, 188, 203, 205, 216, 242, 243 primary body cavities 245 primordial germ cells 267 proboscis 299, 304, 306, 308, 309, 319–321 proboscis skeleton 305, 318, 320, 321 procuticle 94, 95 Prometheus larva 87, 88, 90–93, 95, 97–99, 101, 102, 104, 105 protocoel 286, 287, 304, 306, 314 protoconodont 165 Protoglossus 304–306, 309, 319 Protosagitta 165 protosome 299 Protostomia 111 prototroch 143, 144, 146–148 Pseudicyema 3, 6 Pseudopedicellina 111, 152, 153, 156 Pseudosagitta 165, 175–177, 179, 183, 191, 196 pseudosarcomeres 238 pseudosegment 56, 58, 61, 62, 67 pseudostolons 112 Pterokrohnia 177, 182, 183, 190, 193 Pterosagitta 171, 173, 183, 184, 190 Ptychodera 299, 300, 305, 307, 314, 318 pycnocline 194 pygochord 303, 310 Pyrenomonas 45

Index 

radiation 165 rami 72–74 reaggregation 41 rearrangements 51 regeneration 134, 135 regenerative capacity 41 retractors 68 retrocerebral glands 62 retrocerebral organ 212, 213, 220, 221, 225 RFamide-like neuropeptides 225 RFamide-like peptides 99 RFamidergic 124 rhabdomeres 215 Rhodomonas 45 Rhodope 48 rhombogen 1–4 Rhopalura 11–13, 18, 27–29, 31, 33, 35–37 Rissoa 36 Ritteria 319, 320 rotatory organ 56, 61, 63 Rotifera 55, 62, 83 Saccobdella 83 Saccoglossus 299, 300, 303, 305, 307, 308, 312–315, 319 Sagitta 176, 178, 184, 186, 190, 193, 221, 237, 240, 247, 266 Salinella 1 salinity 177 salivary glands 66, 70 Sangavella 111, 113, 152, 153, 156, 157 sarcomeres 233, 237, 240 Saxipendium 303, 306, 319, 320 Schizocardium 300, 303, 307, 309–311, 315, 318 schizocoely 263–265, 304, 314 Seison 55, 57–59, 61–65, 67, 69, 70, 72–75, 78–84 self-fertilization 8, 261 seminal receptacle 164, 256 seminal vesicle 164, 166, 178, 184, 249, 255, 260 sensitive papillae 125 sensory cells 309 septum 112, 114, 122, 123, 155–157, 170, 201, 225, 229, 243, 250, 251, 256, 265, 305, 306 serotonergic 124, 146 serotonin 26, 99, 225 Serratosagitta 176, 178, 184, 188–190, 245, 247, 259 settlement 314, 315 sexual behavior 35 sexual dimorphism 11 sexual reproduction 47 shell gland 136 shiny spheres 43, 45, 49 silicon 205 Sinusoida 150 Solariella 35, 36 Solidosagitta 175, 183, 184, 189, 191, 196 Spadella 164, 165, 169–171, 173, 179, 181, 188, 192, 193, 199, 201, 205, 207, 208, 211, 216, 218, 223, 227, 228, 231, 233, 234, 237, 238, 240, 250, 253, 255, 256, 262, 264, 266 Spengelia 311, 318 Spengel-stage tornaria 313

spermatids 7, 79, 138, 253 spermatocyte 5, 253 spermatogenesis 139, 261 spermatogonia 5, 8, 77, 79, 138, 139, 253 spermatophore 58, 60, 77–79, 81, 249, 255, 256, 259 spermatophore-forming organ 70, 78 spermatozoa 7, 27–29, 64, 79–81, 104, 136, 138–140, 253, 255, 256, 259, 290 spermioducts 249 spermiogenesis 27, 79, 82 sperm transfer 140, 248 spiral cleavage 35 Spiralia 11, 107 stalk 299, 305, 307, 321 star-cell complex 112, 113, 119, 121, 122, 155–157 star-shaped muscle 122 Stereobalanus 302, 306, 310, 319 Stoecharthrum 11, 12, 18, 36 stolon 112–115, 123, 132, 133, 135, 156, 157, 283–286 stomochord 299, 300, 305, 309, 318, 320 sucking disk 112 supercontracting muscles 238 supercontraction 240, 241 suspension-feeding 283 swarmer-like spheres 47 swarmers 47 symbionts 108 symbiotic 1 symbiotic bacteria 207 synapsin 99, 228 synapticles 318, 319 synapticulae 311 syncytium 3 teeth 166, 169, 177, 178, 182, 183, 203, 205, 207, 233 Tegella 151 Tergivelum 299, 320 terminal cell 102, 104 terminal organ 76, 133, 134 terminal syncytium 76 tetraneury 150 Tetranychus 261 tetrodotoxin 163 thermocline 188, 194, 195 tonofilaments 115, 116, 123, 199, 201, 203, 234, 260 tornaria 309, 314, 315, 318, 319 Torquarator 320 toxin 207 transplantation experiments 46 Triticella 108 trochophore 150 trophic levels 164 trophi 55, 70, 74, 83, 84 tropical submergence 189 trunk coelom 286, 305, 312 tube building 303 Tubifex 261 ultrafiltration 133, 247 unci 72, 73

 331

332 

 Index

upwelling 194 upwelling area 188 Urnatella 111, 113, 114, 152, 153, 155, 156

vestibular ridge 169 Vibrio 207 vitellogenesis 140, 243

valve 117 vasalike protein 261 venom 205 venom glands 205 vermiform embryo 4 vermiform stages 1, 2 vertical migrations 194 vestibular groove 128, 129, 131 vestibular organs 205 vestibular papillae 169 vestibular pit 169

Willeyia 311, 318 Xenokrohnia 180, 191, 207, 238, 240 Xenopleura 319 Yoda 299, 300, 320, 321 zinc 205 Zonosagitta 171, 178, 185, 186, 190, 191, 193 zooplankters 163 zygote 259, 261